Ecology and
Classification of
North American
Freshwater
Invertebrates
Second Edition
This Page Intentionally Left Blank
Ecology and
Classification of
North American
Freshwater
Invertebrates
Second Edition
Edited by
JAMES H. THORP
Bayard and Virginia Clarkson Professor of Biology
Department of Biology
Clarkson University
Potsdam, New York
and
ALAN P. COVICH
Department of Fishery and Wildlife Biology
Colorado State University
Fort Collins, Colorado
San Diego San Francisco New York Boston London Sydney Tokyo
Front cover images: From Jurine, L. (1820). Histoire des Monocles, qui se trouvent
aux environs de Geneve. J. J. Paschoud, Paris, France.
This book is printed on acid-free paper.
Copyright © 2001, 1991 by ACADEMIC PRESS
All Rights Reserved.
No part of this publication may be reproduced o transmitted in any form or by any
means, electronic or mechanical, including photocopy, recording, or any information
storage and retrieval system, without permission in writing from the publisher.
Requests for permission to make copies of any part of the work should be mailed to:
permissions Department, Harcourt Inc., 6277 Sea Harbor Drive,
Orlando, Florida 32887-6777
Academic Press
A Harcourt Science and Technology Company
525 B Street, Suite 1900, San Diego, California 92101-4495, USA
http://www.academicpress.com
Academic Press
Harcourt Place, 32 Jamestown Road, London NW1 7BY, UK
http://www.academicpress.com
Library of Congress Catalog Card Number: 00-109117
International Standard Book Number: 0-12-690647-5
PRINTED IN THE UNITED STATES OF AMERICA
00 01 02 03 04 05 06 EB 9 8 7 6 5 4 3 2 1
“For aminals and callapiters and other wonders
of nature seen through our children’s eyes.”
Dr. James Thorp
“To my family and to Professor G. E. Hutchinson
for their inspiration and patience.”
Dr. Alan Covich
This Page Intentionally Left Blank
Contents
Contributors xiii
Preface xv
2
AN OVERVIEW OF FRESHWATER
HABITATS
James H. Thorp and Alan P. Covich
I.
II.
III.
IV.
1
INTRODUCTION TO FRESHWATER
INVERTEBRATES
James H. Thorp and Alan P. Covich
I. Introduction 1
II. Approches to Taxonomic Classification 1
III. Synopses of the North
American Freshwater Invertebrates 4
IV. Taxonomic Key to Major Taxa
of Freshwater Invertebrates 16
Literature Cited 18
Introduction 19
Lotic Environments 19
Underground Aquatic Habitats 30
Lentic Ecosystems 33
Literature Cited 39
3
PROTOZOA
William D. Taylor and Robert W. Sanders
I. Introduction 43
II. Anatomy and Physiology 45
vii
viii
Contents
III. Ecology and Evolution 58
IV. Collecting, Rearing, and Preparation
for Identification 69
V. Identification of Protozoa 70
Literature Cited 89
4
PORIFERA
Thomas M. Frost, Henry M. Reiswig,
and Anthony Ricciardi
I.
II.
III.
IV.
Introduction 97
Structure and Physiology 97
Ecology and Evolution 105
Collecting, Rearing, and Preparing Sponges
for Identification 115
V. Classification 116
Literature Cited 130
VI. Identification of North American Genera
of Freshwater Turbellaria 163
NEMERTEA
VII. General Characteristics, External and Internal
Anatomical Features 173
VIII. Ecology 174
IX. Current and Future Research Problems 176
X. Collection, Culturing, and Preservation 176
XI. Taxonomic Key to Species of Freshwater
Nemertea in North America 176
Literature Cited 176
7
GASTROTRICHA
David Strayer and William D. Hummon
I.
II.
III.
IV.
Introduction 181
Anatomy and Physiology 182
Ecology and Evolution 183
Collecting, Rearing, and Preparation
for Identification 188
V. Taxonomic Key 189
Literature Cited 192
5
CNIDARIA
Lawrence B. Slobodkin and
Patricia E. Bossert
I. Introduction 135
II. General Biology of Cnidaria 136
III. Descriptive Ecology of Freshwater
Cnidaria 140
IV. Collection and Maintenance
of Freshwater Cnidaria 150
V. Classification of Freshwater
Cnidaria 151
Literature Cited 153
6
FLATWORMS: TURBELLARIA
AND NEMERTEA
8
PHYLUM ROTIFERA
Robert Lee Wallace and Terry W. Snell
I.
II.
III.
IV.
Introduction 195
Anatomy and Physiology 197
Ecology and Evolution 207
Collecting, Rearing, and Preparation
for Identification 230
V. Classification and Systematics 233
Literature Cited 248
9
NEMATODA AND NEMATOMORPHA
Jurek Kolasa
George O. Poinar
TURBELLARIA
I. Introduction: Status in the
Animal Kingdom 156
II. Anatomy and Physiology 156
III. Ecology 158
IV. Current and Future Research Problems 162
V. Collecting, Rearing, and
Identification Techniques 163
NEMATODA
I. Introduction 255
II. Morphology and Physiology 256
III. Development and Life History 260
IV. Ecology 261
V. Collecting and Rearing
Techniques 265
VI. Identification 269
Contents
NEMATOMORPHA
VII. Introduction 280
VIII. Morphology and Physiology 282
IX. Development and Life History 288
X. Sampling 290
XI. Identification 291
Literature Cited 292
BRANCHIOBDELLIDAE
VI. Introduction to Branchiobdellidae 456
VII. Anatomy and Physiology
of Branchiobdellidans 456
VIII. Ecology and Evolution 458
IX. Identification of Branchiobdellidans 459
Literature Cited 461
10
13
MOLLUSCA: GASTROPODA
ANNELIDA: EUHIRUDINEA
AND ACANTHOBDELLIDAE
Kenneth M. Brown
I.
II.
III.
IV.
Introduction 297
Anatomy and Physiology 298
Ecology and Evolution 302
Collecting and Culturing
Freshwater Gastropods 315
V. Identification of Freshwater Gastropods
of North America 315
Literature Cited 325
Ronald W. Davies and Fredric R. Govedich
I. Introduction to Euhirudinea and
Acanthobdellidae 465
II. Taxonomic Status and Characteristics 466
III. Euhirudinea and Acanthobdellidae 468
IV. Ecology 474
V. Collection and Rearing 486
VI. Identification 486
VII. Taxonomic Keys 489
Literature Cited 497
11
MOLLUSCA: BIVALVIA
Robert F. McMahon and Arthur E. Bogan
I.
II.
III.
IV.
Introduction 331
Anatomy and Physiology 333
Ecology and Evolution 354
Collecting, Preparation for Identification,
and Rearing 393
V. Identification of the Freshwater Bivalves
of North America 397
Literature Cited 416
12
14
BRYOZOANS
Timothy S. Wood
I.
II.
III.
IV.
V.
VI.
Introduction 505
Anatomy and Physiology 505
Ecology and Evolution 511
Entoprocta 515
Study Methods 516
Taxonomic Key 517
Literature Cited 523
ANNELIDA: OLIGOCHAETA, INCLUDING
BRANCHIOBDELLIDAE
15
Ralph O. Brinkhurst and Stuart R. Gelder
Diane R. Nelson
OLIGOCHAETA
I. Introduction to Oligochaeta 431
II. Oligochaete Anatomy and Physiology 432
III. Ecology and Evolution of Oligochaeta 438
IV. Collecting, Rearing, and Preparation of
Oligochaetes for Identification 444
V. Taxonomic Keys for Oligochaeta 446
TARDIGRADA
I.
II.
III.
IV.
V.
VI.
Introduction 527
Anatomy 528
Physiology (Latent States) 535
Reproduction and Development 536
Ecology 539
Techniques for Collection, Extraction,
and Microscopy 543
ix
x
Contents
VII. Identification 544
Literature Cited 546
16
WATER MITES (HYDRACHNIDA)
AND OTHER ARACHNIDS
II. Anatomy and Physiology of Crustacea 780
III. Ecology and Evolution
of Selected Crustacea 787
IV. Collecting, Rearing, and Preparation
for Identification 798
V. Classification of Peracarida and Brachiura 799
Literature Cited 800
Ian M. Smith, Bruce P. Smith, and David R. Cook
I. Introduction to Arachnids 551
II. Water Mites (Hydrachnida) 552
III. Other Arachnids in Freshwater Habitats 651
Literature Cited 653
17
DIVERSITY AND CLASSIFICATION
OF INSECTS AND COLLEMBOLA
William L. Hilsenhoff
I.
II.
III.
IV.
V.
Introduction 661
Aquatic Orders of Insects 664
Partially Aquatic Orders of Insects 681
Semiaquatic Collembola — Springtails 699
Identification of the Freshwater Insects
and Collembola 700
Literature Cited 721
20
OSTRACODA
L. Denis Delorme
I.
II.
III.
IV.
V.
VI.
VII.
VIII.
IX.
X.
XI.
XII.
XIII.
XIV.
18
Introduction 811
Anatomy and Physiology 812
Life History 817
Distribution 820
Chemical Habitat 821
Physical Habitat 823
Physiological and Morphological
Adaptations 826
Behavioral Ecology 827
Foraging Relationships 827
Population Regulation 828
Collecting and Rearing Techniques 829
Uses of Freshwater Ostracoda 831
Current and Future Research Problems 832
Classification of Freshwater
Ostracodes 832
Literature Cited 842
AQUATIC INSECT ECOLOGY
Anne E. Hershey and Gary A. Lamberti
I.
II.
III.
IV.
V.
VI.
Introduction 733
Aquatic Insect Communities 733
Aquatic Insect Life Histories 747
Insect-Mediated Processes 751
Secondary Production 761
Effects of Land Use on Aquatic
Insect Communities 763
VII. Role of Disturbance 765
VIII. Aquatic Insects in Biomonitoring Studies 767
IX. Summary 768
Literature Cited 768
19
INTRODUCTION TO THE SUBPHYLUM
CRUSTACEA
Alan P. Covich and James H. Thorp
I. Introduction 777
21
CLADOCERA AND OTHER
BRANCHIOPODA
Stanley I. Dodson and David G. Frey
I. Introduction 850
CLADOCERA
II. Anatomy and Physiology 851
III. Distribution 860
IV. Behavior 862
V. What Cladocerans Eat and What
Eats Cladocerans 866
VI. Population Regulation 867
VII. Functional Role in the Ecosystem 868
VIII. Paleolimnology and Molecular
Phylogeny 868
IX. Toxicology 871
X. Current and Future Research 873
XI. Collecting and Rearing 873
XII. Taxonomic Keys for Cladoceran Genera 875
Contents
OTHER BRANCHIOPODS
XIII. Anatomy and Physiology
of the Other Branchiopods 891
XIV. Ecology of the Other Branchiopods 893
XV. Current and Future Research Problems 899
XVI. Collecting and Rearing Techniques 899
XVII. Taxonomic Keys for Non-Cladoceran Genera
of Branchiopods 899
Literature Cited 904
22
COPEPODA
Craig E. Williamson and Janet W. Reid
I.
II.
III.
IV.
V.
Introduction 915
Anatomy and Physiology 916
Ecology and Evolution 920
Current and Future Research Problems 933
Collecting and Rearing Techniques 934
VI. Identification Techniques 935
Literature Cited 942
23
DECAPODA
H. H. Hobbs III
I.
II.
III.
IV.
V.
Introduction 955
Anatomy and Physiology 955
Ecology and Evolution 964
Current and Future Research Problems 986
Collecting and
Rearing Techniques 989
VI. Identification 991
Literature Cited 993
Glossary 1003
Subject Index 1021
Taxonomic Index 1037
xi
This Page Intentionally Left Blank
Contributors
Numbers in parentheses indicate the pages on which the authors’
contributions begin.
Arthur E. Bogan (331) North Carolina State Museum
of Natural Sciences, Research Laboratory, Raleigh,
North Carolina 27607.
Patricia E. Bossert (135) Science Department,
Northport High School, Northport, New York
11768.
Ralph O. Brinkhurst (431) Hermitage, Tennessee 37076.
Kenneth M. Brown (297) Department of Biological
Sciences, Louisiana State University, Baton Rouge,
Louisiana 70803.
David R. Cook (551) 7836 North Invergordon Place,
Paradise Valley, Arizona 85253.
Alan P. Covich (1, 19, 777) Department of Fishery and
Wildlife Biology, Colorado State University, Fort
Collins, Colorado 80523.
Ronald W. Davies (465) Faculty of Science, Monash
University, Clayton, Victoria 3168, Australia.
L. Denis Delorme (811) National Water Research
Institute, Canada Centre for Inland Waters, Burlington, Ontario, L7R 4A6.
Stanley I. Dodson (849) Department of Zoology, University of Wisconsin, Madison, Wisconsin 53706.
David G. Frey† (849).
Thomas M. Frost† (97).
Stuart R. Gelder (431) Department of Biology, University of Maine at Presque Isle, Presque Isle, Maine
04769.
Fredric R. Govedich (465) Department of Biological
Sciences, Monash University, Clayton, Victoria
3168, Australia.
†
Deceased.
xiii
xiv
Contributors
Anne E. Hershey (733) Department of Biology, University of North Carolina–Greensboro, Greensboro,
North Carolina 27402.
William L. Hilsenhoff (661) Department of Entomology, University of Wisconsin, Madison, Wisconsin
53706.
H. H. Hobbs III (955) Department of Biology, Wittenberg University, Springfield, Ohio 45501.
William D. Hummon (181) Department of Biological
Sciences, Ohio University, Athens, Ohio 45701.
Jurek Kolasa (155) Department of Biology, McMaster
University, Hamilton, Ontario L8S 4K1 Canada.
Gary A. Lamberti (733) Department of Biological
Sciences, University of Notre Dame, Notre Dame,
Indiana 46556.
Robert F. McMahon (331) Department of Biology, The
University of Texas at Arlington, Arlington, Texas
76019.
Diane R. Nelson (527) Department of Biological Sciences, East Tennessee State University, Johnson City,
Tennessee 37614.
George O. Poinar, Jr. (255) Department of Entomology, Oregon State University, Corvallis, Oregon
97331.
Janet W. Reid (915) Department of Invertebrate
Zoology, National Museum of Natural History,
Smithsonian Institution, Washington, DC 20560.
Henry M. Reiswig (97) Redpath Museum and Department of Biology, McGill University, Montreal, Quebec H3A 2K6.
Anthony Ricciardi (97) Department of Biology,
Dalhousie University, Halifax, Nova Scotia B3H
4J1.
Robert W. Sanders (43) Department of Biology, Temple
University, Philadelphia, Pennsylvania 191ZZ.
Lawrence B. Slobodkin (135) Department of Ecology
and Evolution, State University of New York, Stony
Brook, New York 11794.
Bruce P. Smith (551) Biology Department, Ithaca College, Ithaca, New York 14850.
Ian M. Smith (551) Biodiversity Section, ECORC, Central Experimental Farm Agriculture and Agri-Food
Canada, Ottawa, Ontario, Canada K1A 0C6.
Terry W. Snell (195) School of Biology, Georgia Institute of Technology, Atlanta, Georgia 30332.
David Strayer (181) Institute of Ecosystem Studies,
Millbrook, New York 12545.
William D. Taylor (43) Department of Biology, University of Waterloo, Waterloo, Ontario N2L 3G1,
Canada.
James H. Thorp (1, 19, 777) Department of Biology,
Clarkson University, Potsdam, New York 13699.
Robert Lee Wallace (195) Department of Biology,
Ripon College, Ripon, Wisconsin 54971.
Craig E. Williamson (915) Department of Earth and
Environmental Sciences, Lehigh University, Bethlehem, Pennsylvania 18015.
Timothy S. Wood (505) Department of Biological
Sciences, Wright State University, Dayton, Ohio
45435.
Preface
It is a pleasure to write the preface to the second
edition of this book because it indicates that our authors produced a first edition that met the needs of our
audience. To justify having those same professionals
find the second edition useful, as well as attracting a
new generation of readers, it was incumbent upon us
to develop a text that improves and updates the first
edition. We hope that you will find our authors have
done so!
The reader will discover that almost every chapter
has undergone major changes, with updated literature
references, new coverage of invertebrate biology and
ecology, and extensive revisions to taxonomy and
classification. In a few cases, authors have made important improvements in figures and have introduced more
detailed taxonomic keys. We have expanded the
number of authors (for Chapters 4, 11, 13, 16, and 22)
and added a new chapter (Chapter 18 by Drs. Anne
Hershey and Gary Lamberti). As in the first edition,
we limited our coverage of aquatic insects to save
space and avoid excessive duplication with current
textbooks on aquatic insect ecology and classification.
However, we opted to add a chapter on ecology to provide an expanded coverage of aquatic insects at the introductory level to accompany a more taxonomically
centered chapter on insects (Chapter 17 by Dr. William
Hilsenhoff).
This edition is strongly focused on freshwater taxa
found in surface waters of the United States and
Canada. This encompasses at least 10,000 – 15,000
named species while avoiding the broad diversity of
tropical aquatic invertebrates in North America.
Groundwater and estuarine species are typically ignored or barely mentioned. We have also severely
limited coverage of parasitic species and taxa that are
only occasionally associated with aquatic habitats.
As in the first edition, we have asked the authors to
set the taxonomic keys in their chapters at a level
xv
xvi
Preface
where the average reader could reasonably be assured
of reaching an accurate organismal identification without being an expert on that group. This approach is
counter to a tendency in some texts to write dichotomous keys to the species level for the nonexpert. In
many highly specific keys, nonspecialists use the keys to
obtain both a species name and, sometimes, a false
sense of security that accuracy has been achieved. To
supplement information at the generic or higher level in
our text, the chapters provide information on where
the reader can find more detailed, regional taxonomic
keys.
As in our first book, many people contributed to
this edition besides the authors. At Academic Press,
we appreciate, in particular, efforts of our editor, Dr.
Charles R. Crumly, and his able colleague and assistant, Ms. Donna James. At Clarkson University, we especially thank Linda Delosh and Bobbi Baillargeon for
help in managing aspects of the book’s production and
chapter distribution. Each chapter was reviewed by one
to three outside experts; their help is acknowledged in
some chapters and through individual letters from editors and authors. We also express our appreciation
again to the following reviewers of the second edition,
while sincerely apologizing in advance for any we have
overlooked: Drs. Thomas A. Langen, Denis H. Lynn,
Michael L. Pace, Henry M. Reiswig, Anthony Ricciardi, David Strayer, Mitchell J. Weiss, John W. Fleeger,
W. T. Edmondson, Jeffrey D. Jack, Elizabeth J. Walsh,
Andreas Schmidt-Rhaesa, Dreux J. Watermolen, Eileen
H. Jokinen, Charles Lydeard, Caryn C. Vaughn, Mark
J. Wetzel, Dean W. Blinn, Byron T. Backus, Judith E.
Winston, Clark W. Beasley, William R. Miller, John C.
Conroy, Thomas Simmons, Sharon K. Jasper, Manuel
L. Pescador, Arthur C. Benke, Timothy B. Mihuc,
David C. Culver, B. Brandon Curry, Alison J. Smith,
John E. Havel, William R. DeMott, Anna M. Hill, and
David M. Lodge. We continue to appreciate suggestions from readers on how individual chapters or the
book as a whole could be improved. Future comments
and suggestions can be directed to authors of individual chapters or to the editors.
One of the sad tasks not faced in the first edition is
announcing the demise of past authors. We greatly
regret the passing of Professors David G. Frey (a coauthor of Chapter 21: Cladocera and Other Branchiopoda) and, at a relatively young age, Thomas M.
Frost (senior author of Chapter 4: Porifera). Dr. Stanley
I. Dodson handled all revisions to Chapter 21, and
Tom Frost had finished coauthoring Chapter 4 before
he died in a tragic accident in the summer of 2000.
They will be sorely missed by the two of us and by
their many other colleagues and friends.
As premature as it may sound, we have already begun thinking about our third, and probably last edition
of this book with the current editors. While we expect
another ten years will have elapsed before the third edition is completed, we realize it will require new authors
in a number of areas as more senior people retire, shift
research focal areas, or move into academic administration. If you are a reader who has the expertise, interest, and drive to write such a chapter, please feel free to
contact one of us . . . but wait a few years so we have
time to relax and handle those other tasks that have
piled up on our desks while we completed work on the
second edition!
James H. Thorp and Alan P. Covich
1
INTRODUCTION TO FRESHWATER
INVERTEBRATES
James H. Thorp
Alan P. Covich
Department of Biology
Clarkson University
Potsdam, New York 13699
Department of Fishery and Wildlife Biology
Colorado State University
Fort Collins, Colorado 80523
I. Introduction
II. Approaches to Taxonomic Classification
III. Synopses of the North American Freshwater
Invertebrates
A. Protozoa
B. Porifera
C. Cnidaria
D. Flatworms: Turbellaria and
Nemertea
E. Gastrotricha
F. Rotifera
G. Nematoda and Nematomorpha
H. Mollusca
I. Annelida
J. Bryozoa
K. Tardigrada
L. Arthropoda
IV. Taxonomic Key to Major Taxa of
Freshwater Invertebrates
Literature Cited
I. INTRODUCTION
II. APPROACHES TO TAXONOMIC CLASSIFICATION
Next to their more vibrant, and often larger, marine
relatives, freshwater invertebrates may initially seem
drab and uninspiring. Yet once students of freshwater
biology penetrate beyond superficial appearances and
thoroughly examine the diverse structural, physiological, behavioral, and general ecological adaptations of
freshwater invertebrates, few fail to be impressed by the
enticing nature of these fascinating animals.
This chapter serves as a very brief introduction to
freshwater invertebrates of Canada and the United
States. Following a short discussion of procedures for
naming and classifying organisms, we briefly describe
the taxonomic groups covered in Chapters 3 – 23. We
also include a taxonomic key to help you begin the
process of classifying freshwater invertebrates. This
dichotomous key will lead you to the appropriate chapter for more detailed biological and taxonomic information. Additional information on freshwater invertebrates can be found in books by Hutchinson (1993) and
Merritt and Cummins (1996) as well as in various texts
on limnology and stream ecology (e.g., Wetzel, 2001).
No phylum is totally immune to an occasional
reshuffling of species, renaming of constituent taxa, and
recalculation of total species richness (number of
species). This dynamic feature of the discipline is, in
part, a necessary response to the acquisition of new
knowledge, but it also results from the ubiquitous and
almost inherent disagreements that usually typify
groups of two or more scientists! Taxonomists are
frequently labeled as “splitters” or “lumpers” according
to whether, respectively, they tend to acknowledge more
or fewer species within an assemblage. Classifications
they produce based on diverse scientific approaches,
such as cladistics and numerical ( phenetics) taxonomy, can produce dramatically different views of relationships among taxa. Moreover, molecular systematics
has sometimes shaken the traditional taxonomic
schemes based on classical phenotypic characters.
Although arguments at the species level are especially
rampant, debates occasionally break out regarding the
categorization of higher tiers, including classes and
phyla. While reading through this book, you will often
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
1
2
J. H. Thorp and A. P. Covich
encounter references to current taxonomic debates. The
final classification of taxa has been left to the authors of
individual chapters. As editors, we have allowed the
authors a great amount of leeway in this area, requesting only that they inform the reader about the principal
areas of dispute.
Species are named today using a system of binominal nomenclature ( names composed of two parts)
based on the 18th-century proposal of the brilliant
Swedish naturalist Carl von Linné (or Carolus Linnaeus, the latinized form he preferred to use). This is
the “genus and species” designation familiar to most
biology students (e.g., Homo sapiens for the human
species). You will occasionally see a species designated
with three names (a trinomen), which is permissible under guidelines of an international code described below.
For example, Bosmina (Sinobosmina) freyi is the genus,
subgenus, and species name of a water flea, or cladoceran, common in rivers (it is one of a species complex
formerly called Bosmina longirostris). In contrast,
Pheidole xerophila tucsonica is the genus, species, and
subspecies name of a harvester ant living in the desert
near Tucson, Arizona. Scientific names are either in
Latin or in a form that has been latinized (e.g., B.(S.)
freyi was formed by latinizing the last name of
Dr. David G. Frey, to honor this now-deceased coauthor of Chapter 21).
The final arbiter for naming taxa according to this
system is the International Commission on Zoological
Nomenclature, a judicial body elected periodically by
the International Congress of Zoology to evaluate taxonomic names proposed by scientists. They use a set of
agreed-upon rules, termed the International Code of
Zoological Nomenclature (ICZN). The commission
bases its decisions on rules covering the priority of
names and designation of “type” taxa. As the name implies, “priority” specifies that the first name assigned in
a publication is generally the official name of that
taxon. The commission has, however, plenary powers
to set aside the “rightful” name of a species if doing so
would further the cause of science. For example, alteration of a familiar, long-standing name could cause major confusion in the scientific literature. Occasionally,
two distinct taxa are inadvertently assigned the same
binominal name by scientists working in different institutions. In that case, the species described last loses its
name as a junior homonym. To lessen the state of confusion and diminish possibilities for future arguments,
one individual of each species is designated as the
prime example of that taxon. Such “type” specimens
are deposited in one of many internationally recognized
museums and made available for study by responsible
scientists. Type specimens can be replaced only if the
original is inadvertently lost or destroyed.
Taxonomic arguments among scientists fall into
two broad categories: “What is a species?” and “How
do you properly recognize evolutionary relationships
among species?” In the first case, phylogenetic (including are they interfertile), evolutionary, and recognition
definitions are employed. In the second instance, scientists use phenetics, cladistics, and phylogenetic taxonomy to establish relationships, as described below.
Systematists, which include many authors of our
text, are involved in naming species (alpha taxonomy)
and establishing phylogenetic relationships among
groups (beta and gamma taxonomy). Most of these scientists can be divided in modern times into those employing numerical taxonomy ( numerical phenetics)
and those using the more popular cladistics ( phylogenetic systematics), although there is unacknowledged
blending of the two approaches in some cases (especially when gene sequencing data are being used). Each
approach analyzes attributes of a species, or “characters,” (e.g., morphological features or genetic sequences), to determine relationships among groups. In
a very simplified definition, numerical taxonomists sum
a large number of randomly selected characters possessed by multiple species and determine phylogenetic
relationships based on the degree of similarity among
species (i.e., shared characters), without placing relative
evolutionary values on different characters. In contrast,
cladists emphasize the recency of common descent
(shared “derived” characters), a genealogical technique
that places relative evolutionary values on different
characters. Their approach depends significantly on
identification of primitive ( ancestral) vs derived (i.e.,
more recently evolved) characters.
Information on systematic zoology is published in
various journals, such as the Zoological Record,
Zeitschrift fv̈r Wissenschaftliche Zoologie (Abteilung
B), and Bulletin Signaletique (CNRS), as well as in
numerous society publications and journals dealing
with a broad range of subjects including classification.
Readers interested in knowing where best to search for
papers on a given taxon should consult the Literature
Cited section in the appropriate chapter of this book,
or see more specialized texts, such as Merritt and
Cummins (1996) on genera of freshwater insects.
The International Code of Zoological Nomenclature passes judgment on names of genera and species
but does not strictly control higher classification. For
that reason, some disagreements exist about the numbers and names of higher classification tiers, especially
at the ordinal level and above (e.g., the phyla Cnidaria
vs Coelenterata, Bryozoa vs Ectoprocta, and disagreements on whether Crustacea is a phylum, subphylum,
or class). The system for adding suffixes to taxonomic
names as a means of designating classification level has
1. Indroduction to Freshwater Invertebrates
TABLE I Principal Zoological Ranks and Suffixes
Where Designated
Kingdom
Phylum
Subphylum
Superclass
Class
Subclass
Cohort
Superorder
Order
Superfamily (-oidea)a
Family (-idae)
Subfamily (-inae)
Tribe (-ini)a
Subtribe (-ina)b
Genus
Subgenus
Species
Subspecies
a
An ending recommended but not mandatory according
to the International Code of Zoological Nomenclature.
b
An ending customary but not cited in the code.
been confusing in the past but is beginning to change
for the better. A nonexclusive list of the principal zoological ranks and suffixes is shown in Table I.
TABLE II
3
The importance of correct identification and consistent approaches to classification soon becomes apparent
to all people interested in learning about general biotic
relationships. Vast amounts of information are catalogued in research libraries and museums that use the
Linnean system of binominal classification. Once a specimen is correctly identified, a large number of pertinent
references in specialized journals can be accessed, not
only in systematics but also in ecology, evolution, animal
behavior, and medical sciences. Furthermore, the global
revolution known as the Internet has opened a tremendous resource for taxonomists and other biologists. By
using computer-based information retrieval systems present in most libraries or your personal access to the
World Wide Web, you can rapidly track down biological
and taxonomic research on an incredible number of
species. The ease with which student and advanced
scientists can now obtain information on many groups
was almost unimaginable 10 years ago. To help you get
started, we have included in Table II some of the favorite
sites recommended by our authors. Keep in mind, however, that these are only a small minority of available
sites and that the life span of a particular Web address is
unpredictable and relatively fleeting compared to many
of the sources used by investigators in the past (unless
the Web site is directly maintained by a museum).
Brief Selection of Web Sites for Freshwater Invertebrates
Taxonomic group and site
General sitesa
Aquatic Sciences and Fisheries Abstracts
Cambridge Biological Abstracts
Current Contents
Science Citation Index
Web of Science
Zoological Record
Invertebrate Classification and Ecology in Generalb
benthos.org /
nysfola.org / links.html
ucmp.berkeley.edu / help / taxaform.html
york.biosis.org / zrdocs / zoolinfo / grp_dipt.htm
Protozoa
uga.edu / protozoa / home.htm
130.158.208.53 / WWW / Servers / Protistologists.html
megasun.bch.umontreal.ca / protists / protists.html
Porifera
mailbase.ac.uk / lists / porifera / files /
ucmp.berkeley.edu / porifera / porifera.html
Rotifera
member.aol.com / bdelloid1 / deloid.htm
nazca.mmtlc.utep.edu / ~rotifer / main.html
Nematoda and Nematomorpha
ianr.unl.edu / son /
Site maintained by
Corporate
Corporate
Corporate
Corporate
Corporate
Corporate
North American Benthological Society
Individuals
Paleontology Museum, UC Berkeley
BIOSIS (not-for-profit organization) and the Zoological Society of London
Society of Protozoologists
Corporate / University
University of Montreal
Individuals
Paleontology Museum, UC Berkeley
Individuals
Individuals
Society of Nematologists
(Continues)
4
J. H. Thorp and A. P. Covich
TABLE II (Continued)
Taxonomic group and site
Mollusca: Gastropoda
flmnh.ufl.edu / natsci / malacology / malacology.htm#Top
ummz.lsa.umich.edu / mollusks / index.html
Mollusca: Bivalvia
courses.smsu.edu / mcb095f / gallery /
inhs.uiuc.edu / cbd / collections / mollusk.html
Annelida
inhs.uiuc.edu / ~mjwetzel / mjw.inhsCAR.html
keil.ukans.edu / ~worms / annelid.html
Ectoprocta (Bryozoa)
civgeo.rmit.edu.au / bryozoa / default.html
Arthropoda: Insecta and Collembola
afn.org / ~iori / oinlinks.html
entweb.clemson.edu / database / trichopt /
coleopsoc.org /
famu.org / mayfly /
mc.edu / ~stark / stonefly.html
ouc.bc.ca / eesc / iwalker / intpanis /
Arthropoda: Crustacea: Crustacea: General and Peracarida
odu.edu / ~jrh100f / amphome /
york.biosis.org / zrdocs / zoolinfo / arthrop.htm#crustaceans
nmnh.si.edu / gopher-menus / Isopods.html
Arthropoda: Crustacea: Ostracoda
uh.edu / ~rmaddock / IRGO / irgohome.html
Arthropoda: Crustacea: Branchiopoda (Cladocera)
cladocera.uoguelph.ca /
Arthropoda: Crustacea: Copepoda
nmnh.si.edu / iz / copepod /
Arthropoda: Crustacea: Decapoda
bioag.byu.edu / mlbean / crayfish / crayhome.htm
nmnh.si.edu / gopher-menus / Crayfish.html
Site maintained by
Florida Museum of Natural History
Museum of Zoology, University Michigan
Individuals
Illinois Natural History Survey
Illinois Natural History Survey
Individuals / University of Kansas
Individuals / Royal Melbourne Institute of Technology
Individuals
International Trichopteran Committee
Coleopterists Society
Florida Agriculture & Mechanical University
Individuals
Individuals
Individuals
BIOSIS
National Museum of Natural History
International Research Group on Ostracoda, University Houston
Individuals
National Museum of Natural History
Monte Bean Life Science Museum, BYU
National Museum of Natural History
Note. Most of these sites have links to many other sites.
a
We do not guarantee the accuracy or quality of any of these sites. All of these sites were open in the spring of 2000, but we have no idea how
long they will remain accessible. Sites maintained by museums are likely to be long-lasting. Sites designated as being maintained by “individuals” are usually maintained by someone at a university, but the commitment of that university to the long-term survival of the site is usually
unknown.
b
These require that your institution subscribe to the service (i.e., someone pays!).
c
Many of these Internet Web sites begin with http: / /www. Delete the www if those letters do not work.
III. SYNOPSES OF THE NORTH AMERICAN
FRESHWATER INVERTEBRATES
All major phyla of invertebrates, other than Echinodermata, have some freshwater representatives, but
only a few are more diverse in freshwaters than in the
world’s oceans. Some salient features of these freshwater taxa, as discussed in Chapters 3 – 23, are summarized in the remainder of this section.
A. Protozoa
Protozoa are unicellular (or acellular) eukaryotes
existing either as individuals or as members of a
loose-knit colony (Fig. 1). The term protozoa is a
taxon of convenience, having no significance in classification schemes within the kingdom Protista (or
Protoctista) other than as a traditional functional
grouping of all heterotrophic and motile species. Modern taxonomic treatments divide the kingdom Protista
into 27 phyla (more or less), 2 of which contain significant numbers of free-living, phagotrophic species.
These two phyla, Ciliophora (ciliates) and Sarcomastigophora (the majority of amoeboid protozoa),
are the subject of Chapter 3, by William D. Taylor
and Robert W. Sanders.
Protozoa play important, though often overlooked,
roles in freshwater ecosystems. For example, flagellate
protozoans are the predominant predators of picoplankton (primarily bacteria and small cyanobacteria)
1. Indroduction to Freshwater Invertebrates
5
of canals permeating the entire sponge body. Water is
propelled through this canal system both passively, as a
result of its basic hydrodynamic structure, and actively
by choanocytes (collar cells). These cells also extract
bacteria and algae from the water column by suspension (filter) feeding. Many species also contain symbiotic algae (green algae, except for one sponge that
shelters a yellow-green algal symbiont), which provide
supplemental nutrition.
The annual life cycle of a freshwater sponge is
characterized by shifts between periods of active
growth and dormancy in the gemmule stage. Availability of suitable hard substrate and adequate food seem
to be the critical factors regulating the spread and
growth of sponges, as described by Thomas M. Frost,
Henry M. Reiswig, and Anthony Ricciardi in Chapter
4. Few animals prey on sponges, and most of those
graze only a small portion of the sponge’s tissues, leaving the sponge alive to grow and reproduce.
FIGURE 1 Solitary and colonial protozoa: (A) The zooflagellate
Khawkinea; (B) Amoeba; (C) the colonial ciliate Vorticella; and (D)
the solitary ciliate Stentor. See Chapter 3. [These and subsequent figures in this chapter were redrawn and modified from illustration in
Chapters 3 – 22.]
and thus are essential components of the “microbial
loop” within lakes and rivers. Many genera consume
bacteria within the surficial layer of sediments and
in the water column, and larger taxa capture and
eat algae or other protozoa. They, in turn, are preyed
upon by some species of oligochaetes, chironomids,
and rotifers.
B. Porifera
About 27 species of sponges colonize freshwater
habitats in Canada and the United States, with a general east-to-west and north-to-south decline in species
richness. Although less diverse than protozoa, freshwater members of the phylum Porifera — the sponges —
nonetheless are commonly found on hard substrates in
many lakes and streams (Fig. 2). Except for the marine
mesozoa and placozoa, sponges have the simplest
structure of all multicellular phyla, showing little variation in external form and having a cellular-level (or incipient, tissue-level) organization. The most prominent
feature of their anatomy is the progressively finer series
FIGURE 2
Freshwater sponges (phylum Porifera): (A) Drawing of a
living sponge on a stick; (B) representative sponge spicules (all
25 – 150 m in length). See Chapter 4.
6
J. H. Thorp and A. P. Covich
FIGURE 3
Freshwater coelenterates (phylum Cnidaria): (A) A polypoid Hydra; (B) a medusoid Craspedacusta. See Chapter 5.
C. Cnidaria
Cnidarians, or coelenterates, have been relatively
unsuccessful in adapting to freshwater, being represented only by one class (Hydrozoa) with four or five
purely freshwater genera: the common hydras (brown
Hydra and green Chlorohydra, or H. viridissima) and
the rare medusoid Craspedacusta and polypoid Calposoma and Polypodium (Fig. 3). Cordylophora, a mostly
estuarine taxon, occasionally penetrates relatively freshwaters, where it lives in a colonial, sessile form. Despite
their low diversity, cnidarians — or at least hydras — are
routinely present on almost any reasonably hard substrate in permanent ponds and lakes and in streams
ranging from headwater streams to large rivers.
The cnidarian body consists of a generally thin,
acellular layer of mesoglea sandwiched between two
thicker cellular layers (ecto- and endoderm), all surrounding a central body cavity (the coelenteron). Coelenterates dispatch their mostly invertebrate prey
(both zooplankton and some benthos) with ectodermal
batteries of stinging or sticky nematocysts. They are, in
turn, fed upon by a few flatworms and sometimes crayfish. Green hydra are named for the presence of endosymbiotic Chlorella, algae that provide a significant
source of nutrition for cnidarians. The biology of freshwater Cnidaria is described in Chapter 5, by Lawrence
B. Slobodkin and Patricia E. Bossert.
more than 200 species of turbellarians (phylum Platyhelminthes) and three species of ribbon worms (phylum
Nemertea, or Rhynchocoela) found in North America
(Fig. 4). He discusses the acoelomate Turbellaria, the
simplest bilaterally symmetrical eumetazoan with three
embryonic cell layers, and the more complex, coelomate Nemertea, which has an anus and a closed
circulatory system. These latter flatworms possess an
D. Flatworms: Turbellaria and Nemertea
In Chapter 6 on unsegmented flatworms, Jurek
Kolasa describes the biology and classification of the
FIGURE 4 Turbellarian flatworms (phylum Platyhelminthes) and a
ribbon worm (phylum Nemertea): (A) Macrostomum; (B) Dugesia;
and (C) the numertean Prosotoma. See Chapter 6.
1. Indroduction to Freshwater Invertebrates
eversible, muscular proboscis resting in a rhynchocoel.
Despite their common name, flatworms are usually
cylindrical in cross section!
Flatworms are well represented in the benthos of
lakes and streams. Ribbon worms of the single North
American genus Prostoma are rarely reported but can
be common in their typical shallow, littoral zone habitats; all are carnivores that capture their prey with a
sticky proboscis. The much more diverse turbellarians
are distributed more widely than ribbon worms; but
they are not much better known, in part because they
are very difficult to identify with the collection and
preservation techniques normally employed in field
studies. The great majority of these flattened or cylindrical worms are free-living denizens of lakes and
streams from shallow to deep waters. Most are small
(1 mm long), especially the more diverse microturbellaria (~150 species in North America). Better known,
however, are the larger triclads (most 10 mm long),
which are often called planaria. Turbellarians consume
prey in accordance with their size, variously eating bacteria, algae, protozoa, and small or large invertebrates.
E. Gastrotricha
The pseudocoelomate gastrotrichs (Fig. 5) are common residents of epigean waters throughout North
America, with densities ranging from 100,000 to
1,000,000 animals per square meter. Although fewer
than 100 freshwater species from North America
have been reported, the actual diversity of the phylum
7
Gastrotricha is probably much greater in freshwater
(and many marine species are also present). Possibly
because gastrotrichs are so small (50 – 800 m), our
ecological knowledge of this group has not advanced
very far. They colonize both aerobic and anoxic surface
sediments, thrive as members of the aufwuchs assemblage on aquatic plants, and occasionally venture into
the plankton. Oddly enough, given their size, they are
very rare in groundwater (other than the hyporheic
zone of streams). Gastrotrichs thrive on a diet of bacteria, algae, protozoa, and detritus. Not much is known
about their enemies, but these certainly include
amoeba, cnidarians, and predaceous midges. Their
ecology and classification are examined in Chapter 7
by David L. Strayer and William D. Hummon.
F. Rotifera
Perhaps no other phylum is as clearly associated
with freshwater as is Rotifera (Fig. 6). Nearly 2000
species of rotifers, or “wheel animals,” inhabit freshwaters throughout the world, whereas only about 50
species are exclusively marine. These unsegmented,
pseudocoelomates are distinguished by two principal
anatomical features: an apical, ciliated region known as
the corona and a muscular pharynx, termed the mastax, with its complex set of hard jaws. The ciliated
corona is employed for both locomotion and foodgathering. Rotifers range in size from minute creatures
barely 100 μm long to giants of 2 mm or more!
As Robert L. Wallace and Terry W. Snell point out
in Chapter 8, rotifers are one of the three principal animal taxa in the plankton (along with protozoa and microcrustaceans). Because they are more efficient than
cladocera when feeding on minute algae, rotifers can
exert a greater grazing pressure on the small picoplankton. Other rotifers are important predators on bacteria,
protozoa, and small metazoa in the plankton. Rotifers
play a critical role in the microbial (nutrient) loop
within freshwater lakes and rivers. Rotifers are the numerical dominants of most large river zooplankton
communities. Many species are also benthic or nearly
so. Larval fish, some protozoa, insect larvae, microcrustaceans, and other rotifers are numbered among
their predators.
G. Nematoda and Nematomorpha
FIGURE 5 Representative freshwater Gastrotricha: (A) Chaetonotus;
(B) Stylochaeta. See Chapter 7.
Nematodes, which are best known to the average
person for damaging crops, are also abundant in freshwater lakes and streams where they spend all or a
large part of their life cycle (Fig. 7A). Chapter 9 by
George O. Poinar covers 66 genera (encompassing
300 – 500 species), representing over 95% of the
8
J. H. Thorp and A. P. Covich
FIGURE 6 “Wheel animals” of the phylum Rotifera: (A) A Solitary Keratella; (B) a colony
of Sinantherina. See Chapter 8.
known species of roundworms. Nematodes are unsegmented worms with a body cavity (pseudocoel) and
complete alimentary tract; they lack jointed appendages and specialized circulatory and respiratory
systems. Roundworms are significant components of
the micro- and macrobenthos of many aquatic ecosystems. For example, in Mirror Lake, New Hampshire,
they constitute 60% of all individuals in the benthic
metazoa, although only about 1% of the total biomass
(Strayer, 1985, as cited in Chapter 9). Many nematodes are parasitic in plants and animals, but these are
only briefly mentioned in Chapter 9 by George O.
Poinar. Free-living roundworms, the focus of that
chapter, obtain nourishment from bacteria, algae, and
protozoa. While flatworms and crayfish feed occasionally on roundworms, their greatest threat comes from
other, predaceous nematodes.
Another phylum of worms that, in a very
rough way, resembles nematodes is Nematomorpha
(Fig. 7B). These hairworms, or horsehair worms, are
free-living as adults but parasitic as larvae in various
invertebrates. Like nematodes, they are unsegmented
pseudocoelomates, but their intestine is nonfunctional,
and they are generally much larger than roundworms.
The evolutionary position of nematomorphs remains
controversial, but they probably have a distant relationship with nematodes. All but one genus occurs in
freshwater, and seven genera from North American
lakes and streams are reported.
H. Mollusca
Two classes within the phylum Mollusca contain
large numbers of freshwater species: the classes Gastropoda (snails) and Bivalvia (mussels and clams).
These ubiquitous and abundant molluscs are discussed
separately in Chapters 10 (snails) and 11 (mussels and
clams).
1. Gastropoda
FIGURE 7 Illustrated is a common roundworm (A; phylum
Nematoda) and a less frequently observed, adult horsehair worm (B;
phylum Nematomorpha). See Chapter 9.
Snails are one of the more diverse freshwater
groups (having about 659 species in North America)
and are certainly among the most easily recognized by
the public. These soft-bodied, unsegmented coelomates
have a body organized into a muscular foot, distinct
head region, and visceral mass: a fleshy mantle covers
the viscera and secretes a spiraled, univalve, calcareous
shell (Fig. 8). They are commonly found in littoral regions of lentic and lotic habitats, where they collect
bottom detritus, graze on the algal film covering rocks,
wood snags, and plants or even float upside down at
the water surface, consuming trapped and neustonic algae. Recent ecological work on freshwater gastropods,
1. Indroduction to Freshwater Invertebrates
9
species are listed on state and federal lists of “threatened and endangered species” as a result of human
disruption of their aquatic habitats, loss of fish hosts
for larval stage, and invasion of Asian clams (Corbicula fluminea), zebra mussels (Dreissena polymorpha),
and quagga mussels (D. bugensis). These exotic
species have become the numerical and/or biomass
dominants in many North American waters, especially
east of the great plains, and they are exerting substantial ecological and economic effects.
I. Annelida
FIGURE 8 Three types of shell morphologies in freshwater snails
and limpets (phylum Mollusca, class Gastropoda): (A) The most
common, spiral shell, as shown by Pomacea: (B) two views of
the planospiral snail Planorbella (Helisoma); and (C) the conical
freshwater limpet Ferrissia. See Chapter 10.
as described by Kenneth M. Brown in Chapter 10, has
focused on their important roles as grazers on periphyton and as prey of benthic-feeding fish and invertebrates, especially crayfish but also others like leeches
and predaceous water bugs (Belostomatidae). Freshwater limpets are included in Gastropoda, although they
are not related closely to their marine relatives.
The phylum Annelida consists primarily of three
relatively diverse freshwater groups (oligochaete
worms, branchiobdellidans, and leeches) and a few minor freshwater taxa. Annelids are legless, segmented
coelomates with serially arranged organs; they occur in
both lentic and lotic environments, where they tend to
be either detritivores (oligochaetes) or predators
(leeches and branchiobdellidans). The taxonomy of Annelida is in great dispute — an observation reflected by
the diverse views of authors in Chapters 12 and 13. A
strong area of controversy involves the rank of higher
groups, beginning at whether Clitellata (which encompasses all annelids except polychaetes) is a superclass,
class, or subclass; this then affects ranks of lower
groups like the oligochaetes and leeches. Unless the
reader is a student of annelid biology and evolution,
2. Bivalvia
The North American fauna of freshwater bivalves
is the richest in the world, with over 250 species of
mussels and clams (Fig. 9; see also Chapter 11, by
Robert F. McMahon and Arthur E. Bogan). Bivalves,
or pelecypods, are major deposit and suspension feeders within permanent lakes, streams, and large rivers,
where they are often the largest invertebrates in body
mass. Except for the smaller pea clams (Sphaeriidae),
native mussels (Unionacea) are uniquely capable of
growing large enough to escape predation by almost
all aquatic vertebrates and invertebrates. Bivalves are
much less motile than gastropods, and their body
is enclosed in two hinged, calcareous shell valves.
A radula, which is employed by snails for grazing, is
absent in clams and mussels. Unlike the vast majority
of North American freshwater invertebrates, bivalves have been of direct economic importance to humans; they were formerly used for food (by Native
Americans) and button production and are now harvested for freshwater pearls or for the core elements
needed to grow marine-cultured pearls. Many native
FIGURE 9 One native and two invadere species of freshwater
bivalves (phylum Mollusca,class Bivalvia): (A) The native mussel
Quadrula; (B) the ubiquitous Asiatic clam, Corbicula; and (C) a
newcomer to North America, the rapidly spreading zebra mussel,
Dreissena. See Chaper 11.
10
J. H. Thorp and A. P. Covich
one can set aside these arguments for the time being
and concentrate on learning other details of annelid
biology, ecology, and classification.
1. Oligochaeta and Branchiobdellida
In Chapter 12, Ralph O. Brinkhurst and Stuart R.
Gelder discuss the free-living oligochaete worms (~150
species) and the related branchiobdellidans, a lesswell-studied group of 107 species in North America
(Fig. 10). Oligochaetes are among the more widely distributed freshwater taxa and one of the very few invertebrates to colonize moist terrestrial environments. In
addition to the characteristics of the phylum as a
whole, oligochaetes lack suckers and generally have
four bundles of setae on every segment but the first.
Oligochaetes are best known for their ingestion of sediments in the littoral and profundal zones (a process
prominent in nutrient recycling), but some species feed
on benthic algae and other epiphytic organisms. Their
great abundance and ease of consumption and assimilation make them favorite prey items for many invertebrate and vertebrate predators. Branchiobdellidans are
ectocommensals of mostly astacoidean crayfish, where
they consume epibionts living on crayfish, spare food
lost by the host in the feeding process, tissue from host
wounds, and other branchiobdellidans. They have a
posterior sucker for attaching to their hosts.
2. Leeches and Acanthobdellids
Another ecologically important group of annelids
is represented by 73 North American species of leeches,
FIGURE 11
Schematic view of the leech Placobdella. See
Chapter 13.
as described in Chapter 13, by Ronald W. Davies and
Fredric R. Govedich. A single species of leech-like
Acanthobdellidae occurs in North America; it is a temporary ectoparasite of salmonid fish. Unlike most
oligochaetes, leeches have suckers (anterior and posterior) and lack setae (Fig. 11). Another major distinction
between these two annelids is their feeding niches.
Most leeches are predators of invertebrates, such as
chironomid midges, oligochaetes, amphipods, and molluscs; but nonscientists best know this group for the
minority that are ectoparasites, feeding on the blood
of vertebrates (sanguivory), including incautious humans. Predators of leeches include fish, birds, snakes,
salamanders, and a large variety of freshwater invertebrates.
J. Bryozoa
FIGURE 10 Two freshwater annelids with distinctive lifestyles:
(A) The free-living oligochaete Branchiura, showing its posterior
gill filaments; (B) Ceratodrilus, a branchiobdellidan which is an
ectocommensal of crayfish. See Chapter 12.
Bryozoans, or “moss animals,” are generally sessile, colonial invertebrates with ciliated tentacles (called
“lophophore”) for capturing suspended solitary algae,
protozoa, and organic particles (Fig. 12). All but one
North American species are in the phylum Ectoprocta,
or Bryozoa, and are composed of animals whose
mouth, but not anus, is located within the whorl of
feeding tentacles. Although the diversity of freshwater
Bryozoa is not high (24 species in North America), they
are actually quite common occupants of hard, relatively stationary, and biologically inactive substrates.
Most live in relatively warm water (18 – 28°C). They
occur in lakes but thrive better in flowing water, where
1. Indroduction to Freshwater Invertebrates
11
FIGURE 12 Freshwater bryozoans: (A) Plumatella colony (phylum Bryozoa
or Ectoprocta;) (B) close-up of two individuals in an ecotoproct colony; and
(C) the ubiquitous entoproct Urnatella. See Chapter 14.
a constant source of suspended food encounters their
tentacles. Our ecological knowledge of Bryozoa is relatively sparse despite the long-standing familiarity of
scientists with this group. The only nonectoproct
bryozoan is the cosmopolitan, suspension-feeding
Urnatella gracilis. It is classified in the phylum Entoprocta, a taxon with very dubious relation to the
ectoprocts. Both the mouth and anus of entoprocts are
placed within the whorl of feeding tentacles; other important structural differences from the ectoprocts also
exist. The biology, ecology, and classification of
bryozoans are described in Chapter 14, by Timothy
S. Wood.
while 6 genera with 13 – 14 species are typically freshwater. The actual number of water bear species in
North America is probably considerably higher than
this low value. Tardigrades are segmented micrometazoa with four fleshy legs terminating in claws (Fig. 13).
They have piercing/sucking mouthparts and obtain nutrition from plants, animals, and detritus. An important part of the water bear life cycle is the latent stage
(cryptobiosis) entered into when environmental conditions become inhospitable. The biology and systematics
of these little-known creatures are described by Diane
R. Nelson in Chapter 15.
K. Tardigrada
Tardigrades, or “water bears,” are easy to overlook because of their diminutive nature (adults average
250 – 500 m), but they are widely dispersed in freshwater and terrestrial environments, and a few of the
~800 species worldwide occur in the ocean. Although
the taxonomy of Tardigrada needs to be revised, the diversity of freshwater species in North America is low
compared to most other phyla with freshwater members: 3 genera with 5 species are strictly freshwater,
FIGURE 13 Lateral view of a heterotardigrade (phylum Tardigrada).
See Chapter 15.
12
J. H. Thorp and A. P. Covich
L. Arthropoda
The most successful terrestrial phylum and one of
the most prominent freshwater taxa is Arthropoda. Its
three subphyla with freshwater members — Chelicerata
(water mites and aquatic spiders), Uniramia (aquatic
insects), and Crustacea (crayfish, fairy shrimp, copepods, etc.) — are all diverse and important components
of lakes and streams. (Some authors prefer to group
the insects, myriapods, and crustaceans as classes
within a single subphylum, Mandibulata.) Arthropods
occupy every heterotrophic niche in benthic and pelagic
habitats of most permanent and temporary aquatic systems and are well represented in groundwater habitats.
As adults, these metameric coelomates are characterized by a chitinous exoskeleton and stiff, jointed appendages modified as legs, mouth parts, and antennae
(except in water mites).
1. Water Mites
The class Arachnida (subphylum Chelicerata) is
represented in freshwater by a single genus of semiaquatic, true spiders (Dolomedes) and a colossal number of water mites (subclass Acari; Fig. 14). In Chapter
16, Ian M. Smith, David R. Cook, and Bruce P. Smith,
discuss the biology and classification of several distantly related groups of mites, the most diverse of
which is Hydrachnida. As they point out, a single
square meter of substrate from a littoral weed bed in a
eutrophic lake may contain 2000 mites representing up
to 75 species and a comparable area of rocky stream
bed may yield 5000 individuals in 50 or more species!
Over 1500 species are estimated to be present in
Canada and the United States. Because of their prodigious numbers and wide distribution, mites probably
play a major, albeit often ignored, role in aquatic
ecosystems as predators (e.g., on ostracodes, early instars of insects, various zooplankton, and insect eggs),
ectoparasites (mostly on insects), and prey of fish, other
mites, and a few other invertebrates. For example,
FIGURE 14
Generalized larval (A) and adult (B) water mites
(phylum Arthropoda, Acari). See Chapter 16.
mites interact intimately with all life stages of aquatic
insects, parasitizing 20 – 50% of adults in natural populations. Mites differ from members of other arthropod
subphyla in lacking antennae; adults generally have
four pairs of jointed walking legs and segmentation is
greatly reduced or not immediately apparent.
2. Aquatic Insects and Collembola
Perhaps the most diverse and best studied groups
of freshwater invertebrates are the 10 insect orders containing the roughly 6000 species of aquatic insects
(Fig. 15). William L. Hilsenhoff introduces the classification and general characteristics of aquatic insects in
Chapter 17, and Anne E. Hershey and Gary A. Lamberti tackle the daunting task of summarizing some basic ecological aspects of aquatic insects in Chapter 18,
leaving the more detailed discussions to books devoted
to all or specific groups of aquatic insects (e.g., McCafferty, 1981; Ward, 1992; Merritt and Cummins, 1996).
Like the water mites, insects occupy every freshwater
habitat and heterotrophic niche and are present with
tremendous densities and diversities. The majority
spend most of their lives as larvae, only briefly departing the freshwater milieu to mate. Aquatic insects feature mandibles and one pair of antennae; adults have
three pairs of legs and generally possess functional
wings. Chapter 17 also covers the semiaquatic springtails, or Collembola. These wingless arthropods were
once classified as insects but are now generally regarded
as members of the separate class Entognatha within the
superclass Hexapoda, which also includes insects. They
are usually associated with the water surface (neuston)
or riparian habitats. Although a number of species are
sometimes collected in or on water, only about 15
species are regarded as semiaquatic or riparian.
3. Crustaceans
Nearly 4000 species of crustaceans inhabit freshwaters around the world, occupying a great diversity of
habitats and feeding niches. Within pelagic and littoral
zones, water fleas and copepods are the principal
macrozooplankton, and benthic littoral areas shelter
vast numbers of seed shrimps, scuds, and other crustaceans. An omnivorous feeding habit is typical of
crustaceans, although there are many strict herbivores,
carnivores, and detritivores. Members of the subphylum Crustacea are characterized by a head with paired
mandibular jaws, a pair of maxillae, and two pairs of
antennae. Their appendages are often biramous.
a. Peracarida and Branchiura Several groups of
Crustacea, most in the class Malacostraca, superorder
Peracarida, have a relatively small number of freshwater representatives. Two of these, Amphipoda and
1. Indroduction to Freshwater Invertebrates
13
FIGURE 15
Five arthropods in the superclass Hexapoda: (A) Baetis, a mayfly nymph (animals
drawn in a-d are all in the class Insecta); (B) the caddisfly larva Polycentropus; (C) and adult elmid
beetle Stenelmis; (D) the midge larva Chironomus; and (E) a semi-aquatic springtail (class Entognatha,
order Collembola). See Chapter 17.
Isopoda, are important taxa within marine systems.
Chapter 19 (by Alan P. Covich and James H. Thorp)
includes both an introduction to the subphylum Crustacea and coverage of the ecology and systematics of
three malacostracan orders representing less than 200
species: Amphipoda (scuds), Isopoda (aquatic sow
bugs), and Mysidacea (opossum shrimp) (Fig. 16). Of
these three taxa, the scuds are the most widely distributed and diverse within North American freshwaters,
for example. The peracaridans are omnivorous and opportunistic, consuming items ranging from algae and
organic matter to small, living invertebrates and flesh
of dead bivalve molluscs. They occur in ecosystems
ranging in size from large rivers to small ponds and are
also well represented among subterranean fauna. The
subclass Branchiura, or “fishlike,” attach themselves to
the gills of fish; only one genus occurs in North American freshwaters (Argulus).
b. Ostracoda Approximately 420 species of mussel
and seed shrimps (class Ostracoda) found on this continent are easily distinguished from other crustaceans by
the typical, bivalved carapace enveloping their bodies
(Fig. 17). This protective covering is composed of chitin
and calcite. Ostracodes occur within the benthos of
nearly every conceivable aquatic system from temporary
ponds to large rivers to groundwater habitats, but they
rarely enter the plankton even though many species are
good swimmers. Almost all are free living and most are
herbivores on attached algae or detritivores. They are,
in turn, consumed by fish, waterfowl, and various benthic and planktonic invertebrate predators, as discussed
by L. Denis Delorme in Chapter 20.
c. Cladocera and Other Branchiopoda The class
Branchiopoda includes the common Daphnia and other
cladocera frequently studied in the laboratory component of introductory biology courses, along with lesser
known representatives of the class such as the elusive
fairy shrimp of ephemeral pools and saline lakes
(Fig. 18). Branchiopods are a heterogeneous group
linked by similar mouth parts and leaf-like thoracic legs
(phyllopods). Virtually all species within the eight extant
orders of Branchiopoda are limited to freshwaters
(mostly lentic systems). As pointed out by Stanley I.
Dodson in Chapter 21, branchiopods occupy key positions in aquatic communities. As consumers, they are
algivorous herbivores, detritivores (often assimilating
bacteria on benthic or suspended organic matter), and
occasionally predators of small invertebrates. They are
important prey items in the diets of many fish, waterfowl, and certain other vertebrate and invertebrate
predators. Many species are planktonic, while others live
in shallow-water benthic habitats. Some, like the fairy
14
J. H. Thorp and A. P. Covich
FIGURE 16
Representative freshwater crustaceans: (A) Gammarus, a scud (order
Amphipoda); (B) Caecidotea, an aquatic sow bug (order Isopoda); and (C) Mysis, an
opossum shrimp (order Mysidacea). See Chapter 19.
shrimp Artemia, are adapted to temporary ponds and to
hypersaline ponds and lakes. Scientists estimate that the
United States and Canada contain ~400 species of cladocera and 78 species of noncladoceran branchiopods.
d. Copepoda Copepods are very common in
aquatic and semiaquatic habitats, spanning the gamut
FIGURE 17
Female ostracode Candoma (crustacean class Ostracoda) with left value of carepace removed. See Chapter 20.
from ephemeral pools, groundwaters, and wetlands to
the benthos and plankton of large lakes and rivers.
They are also major constituents of marine zooplankton and can be abundant in moist terrestrial environments. Most are free-living, but two of the seven
orders contain species parasitic on freshwater fish.
Slightly over 300 species from North American fresh
waters are described, 10 of which are considered to be
introduced. They typically represent more than 50% of
the zooplankton biomass in rivers and lakes. As was
the case with cladocera, copepods have an integral role
in aquatic food webs both as primary and secondary
consumers (most are omnivorous) and as a major
source of food for many young and adult fish, other
copepods, and other invertebrates, such as larval phantom midges (Chaoborus). Copepods can be distinguished from other small (0.5 — 2.0 mm) crustaceans
by their possession of: (1) a cylindrical, segmented
body; (2) two setose, caudal rami on the posterior end
of the abdomen (which are used in swimming); and
(3) a single, simple anterior eye (Fig. 19). The biology
and classification of this important crustacean are
1. Indroduction to Freshwater Invertebrates
15
FIGURE 18 Tow branchiopod crustaceans: (A) The widely distributed “water flea” Daphnia; (B) the more
localized fairly shrimp Eubranchipus, showing an adult male (C) Triops, a notostracan tadpole shrimp. See
Chapter 21.
discussed by Craig E. Williamson and Janet W. Reid in
Chapter 22.
e. Decapoda Decapoda is the most diverse order
of the class Malacostraca in marine and freshwater
FIGURE 19
ecosystems. It encompasses 342 species of crayfish
and 17 species of shrimp in epigean and subterranean
freshwaters north of Mexico (Fig. 20). All are characterized by terminal claws on the first three of five pairs
of thoracic appendages and a branchial chamber
Two copepod crustaceans: (A) A generalized calanoid species; (B) a
representative cyclopoid copepod. See Chapter 22.
16
J. H. Thorp and A. P. Covich
enclosed within the carapace (see Chapter 22 by
Horton H. Hobbs). Decapods are the largest motile
invertebrates in North American freshwaters. Their
size, activity, and omnivorous diet make them key
players in the benthos of many ecosystems. For example, they can influence species distribution of snails
within a community by selectively preying on certain
size classes or shell morphologies, and they can be
voracious herbivores that devastate vascular plants in
ponds. Decapods, unlike practically all other North
American invertebrates, are increasingly consumed by
humans, especially in the southern United States.
M. Taxonomic Key to Major Taxa of
Freshwater Invertebrates
FIGURE 20 Crayfish (A) are extremely common decapod crustaceans, while palaemonid shrimp (B) have a more restricted distribution (although occasionally abundant). See Chapter 23.
The following dichotomous, taxonomic key is designed specifically to lead the reader to appropriate
chapters within this book (i.e., it should not be employed in other contexts). It can be used for the larval
and adult stages of aquatic insects, but it generally
applies to the adults of other taxa.
1a.
Unicellular (or acellular) organisms present as individuals or colonies; heterotrophic and / or autotrophic; protozoans (Fig. 1)
...............................................................................................................................................................kingdom Protista (Chapter 3)
1b.
Multicellular, heterotrophic organisms (sometimes with symbiotic autotrophs)................................................................................. 2
2a(1b).
Multicellular animals without discrete organs and with a cellular-level (or incipient tissue-level) construction; sponges (Fig. 2)
................................................................................................................................................................phylum Porifera (Chapter 4)
2b.
Multicellular animals with organ or organ-system construction and two or three embryonic cell layers ......................................
.......................................................................................................................................................................................................... 3
3a(2b).
Two embryonic cell layers; adults with a central body cavity opening to the exterior and surrounded by cellular endoderm, and
acellular mesoglea, and a cellular ectoderm; polyps (e.g., hydra) and medusae (e.g., Craspedacusta) (Fig. 3) ................................
...............................................................................................................................................................phylum Cnidaria (Chapter 5)
3b.
Three embryonic cell layers; adults with cellular ectoderm, mesoderm, and endoderm ..................................................................... 4
4a(3b).
Flattened or cylindrical, bilateral, acoelomate worms with only one opening to the digestive tract; flatworms (some occasionally
called planaria) (Fig. 4) ....................................................................................phylum Platyhelminthes: class Turbellaria (Chapter 6)
4b.
Pseudocoelomate or coelomate eumetazoa with a complete digestive tract........................................................................................ 5
5a(4b).
Flattened, unsegmented worms with an eversible proboscis in a rhynchocoel and a closed circulatory system; ribbon worms (Fig. 5)
............................................................................................................................................................. phylum Nemertea (Chapter 6)
5b.
Not with above characteristics ......................................................................................................................................................... 6
6a(5b).
Pseudocoelomates; legs, lophophorate tentacles, and segmentation absent; gastrotrichs (Fig. 5), rotifers (Fig. 6), roundworms
(Nematoda; Fig. 7a), or hairworms (Nematomorpha; Fig. 7b). ......................................................................................................... 7
6b.
Coelomates; legs, lophophorate tentacles, and/or metameres present; snails and mussels (Mollusca; Figs. 8 and 9), segmented worms
(Annelida; Figs. 10 and 11), bryozoans (Fig. 12), water bears (Tardigrada; Fig. 13), and arthropods (mites, insects, and crustaceans;
Figs. 14 – 20) ................................................................................................................................................................................... 10
7a(6a).
Small (50 – 800 m), spindle- or tenpin-shaped, ventrally flattened pseudocoelomates; more or less distinct head bearing sensory
cilia; cuticle usually ornamented with spines or scales of various shapes; posterior of body normally formed into a furca with distal
adhesive tubes; gastrotrichs (Fig. 5) ................................................................................................. phylum Gastrotricha (Chapter 7)
7b.
Not with above characteristics .......................................................................................................................................................... 8
1. Indroduction to Freshwater Invertebrates
17
8a(7b).
Nonvermiform pseudocoelomates with apical (and usually ciliated) corona and muscular pharynx (mastax) with complex set of
jaws; wheel animals, or rotifers (Fig. 6) .................................................................................................. phylum Rotifera (Chapter 8)
8b.
Not with above characteristics; nonsegmented worms ..................................................................................................................... 9
9a(8b).
Alimentary tract present; cylindrical body usually tapering at both ends; noncellular cuticle without cilia, often with striations,
punctuations, minute bristles, etc.; most less than 1 cm long (except family Mermithidae); roundworms (Fig. 7a)........................
........................................................................................................................................................... phylum Nematoda (Chapter 9)
9b.
Degenerate intestine; anterior and posterior tips normally are obtusely rounded or blunt; cuticle opaque and epicuticle usually crisscrossed by minute grooves; pseudocoel frequently filled with mesenchymal cells with large clear vacuoles, giving tissue a foamy appearance; length several centimeters to 1 m, width 0.25 – 3 mm; only adults with free-living stage; hairworms or horsehair worms
(Fig. 7b) ..................................................................................................................................... phylum Nematomorpha (Chapter 9)
10a(6b).
Soft-bodied coelomates, usually with a hard calcareous shell; most with ciliated gills, a ventral muscular foot, and a fleshy mantle
covering internal organs; snails and mussels (Figs. 8 and 9) ............................................................................ phylum Mollusca
........................................................................................................................................................................................................ 11
10b.
Body not enclosed in a single, spiraled shell or in a hinged, bivalved shell, or if a bivalved shell is present, then animal has jointed
legs.................................................................................................................................................................................................. 12
11a(10a).
Body enclosed in a single shell, which usually has obvious spiral coils (limpets are the exception); body basically partitioned into
head, foot, and visceral mass; radula present; snails and limpets (Fig. 8)
............................................................................................................................... phylum Mollusca: class Gastropoda (Chapter 10)
11b.
Body enclosed in two hinged shells; head and radula absent; mussels and clams (Fig. 9)
..................................................................................................................................... phylum Mollusca: class Bivalvia (Chapter 11)
12a(10b).
Legs absent in all life stages ............................................................................................................................................................ 13
12b.
Adults and most larval stages with legs; if larvae without legs or prolegs (some insects), then cephalic region with paired mandibles
.........................................phylum Arthropoda .............................................................................................................................. 16
13a(12a).
Lophophorate tentacles absent; segmented worms . ........................................................................................................................ 14
13b.
Ciliated, lophophorate feeding tentacles present.............................................................................................................................. 15
14a(13a).
No suckers and four bundles of chaetae on each segment except the first (Oligochaeta and Aphanoneura; Fig. 10A) “or” posterior,
disk-shaped sucker on segment 11 and head composed of peristomium and three segments, ectoparasites on crayfish (Branchiobdellida; (Fig. 10B) ....................................................................................................................... phylum Annelida (in part) (Chapter 12)
14b.
Body with anterior and posterior suckers and no chaetae; leeches (Euhirudinea; Fig. 11) “or” posterior sucker on four segments, ectoparasites on salmonid fish (Acanthobdella peledina)........................................................... phylum Annelida (in part) (Chapter 13)
15a(13b).
Anus opens outside a generally U-shaped lophophore; bryozoans (Fig. 12A) ....................................................... phylum Ectoprocta
(Bryozoa) (Chapter 14)
15b.
Both mouth and anus open within lophophore; calyx with single whorl of 8 – 16 ciliated tentacles (Fig. 12b) .................................
........................................................................................................................................................ phylum Entoprocta (Chapter 14)
16a(12b).
Minute adults with four pairs of clawed, nonjointed legs; water bears (Fig. 13) ...........................................................................
......................................................................................................................................................... phylum Tardigarda (Chapter 15)
16b.
Adults and most larvae with jointed legs; arthropods ..................................................................................................................... 17
17a(16b).
Antennae absent; body divided into cephalothorax (prosoma) and abdomen (opisthosoma); adults generally with four pairs of legs;
water mites or the pseudo-aquatic water spiders (Fig. 14) ..........................subphylum Chelicerata, class Arachnida ......................
........................................................................................................................................................................................ (Chapter 16)
17b.
Antennae present; insects, springtails, and crustaceans . .................................................................................................................. 18
18a(17b).
Mandibles and one pair of antennae; adults (most of which are terrestrial) generally with functional wings, larvae with wing pads
or without any indication of wings; larvae and adults normally with three pairs of legs (some larvae without legs); wingless adults in
class Entognatha (order Collembola, the springtails) and winged adults are in class Insecta (aquatic insects) (Fig. 15)
..........................................................................................................................................subphylum Uniramia (Chapters 17 and 18)
18b.
Two pairs of antennae and mandibles (at some life stage); adults with variable numbers of legs, never with wings ............................
....................subphylum Crustacea ................................................................................................................................................. 19
19a(18b).
Body with 19 segments (head: 5; thorax: 8; abdomen: 6); abdomen with pleopods; carapace covers head and all or part of thorax
..............................class Malacostraca ........................................................................................................................................... 20
19b.
Body enveloped in a bivalved carapace (ostracodes), “or” thoracic legs flattened and leaf-like (water fleas and other branchiopods),
“or” 11 postcephalic segments and a single cephalic eye (copepods) .............................................................................................. 21
18
J. H. Thorp and A. P. Covich
20a(19a).
Malacostracans with first three of five pairs of thoracic appendages modified as maxillipeds (including usually large chelae on first
pair in crayfish) and carapace encloses branchial chamber; crayfish and atyid and palaemonid shrimp (Fig. 20)
.............................................................................................................................................................. order Decapoda (Chapter 23)
20b.
Malacostracan crustaceans with other characteristics, including scuds (order Amphipoda), aquatic sow bugs (order Isopoda), and
opossum shrimp (order Mysidacea) (Fig. 16) ................................................................................................................. (Chapter 19)
21a(19b).
Body enveloped in laterally compressed, bivalved carapace; mussel and seed shrimp (Fig. 17) ......................................................
.............................................................................................................................................................. class Ostracoda (Chapter 20)
21b.
Body not enclosed within a bivalved carapace ................................................................................................................................ 22
22a(21b).
Thoracic legs flattened and leaf-like (phyllopod), mandibles are simple unsegmented rods with inner corrugated grinding surfaces;
last body segment with a pair of spines or claws; water fleas (cladocera) and other branchiopods (Fig. 8) ..........................................
......................................................................................................................................................... class Branchiopoda (Chapter 21)
22b.
Cylindrical, segmented body, two setose caudal rami on posterior end of abdomen, and a single, simple anterior eye; copepods
(Fig. 19) ................................................................................................................................................ class Copepoda (Chapter 22)
LITERATURE CITED
Hutchinson, G. E. 1993. A treatise on limnology, Vol. IV, The
zoobenthos. Wiley, New York.
McCafferty, W. P. 1981. Aquatic entomology. Science Books International, Boston, MA.
Merritt, R. W., Cummins K. W., Eds. 1996. An introduction to the
aquatic insects of North America. 3rd ed. Kendall/Hunt,
Dubuque, IA.
Ward, J. V. 1992. Aquatic insect ecology. 1. Biology and habitat.
Wiley, New York.
Wetzel, R. G. 2001. Limnology, 3rd ed. Academic Press, San Diego, CA.
2
AN OVERVIEW OF
FRESHWATER HABITATS
James H. Thorp
Alan P. Covich
Department of Biology
Clarkson University
Potsdam, New York 13699
Department of Fishery and Wildlife Biology
Colorado State University
Fort Collins, Colorado 80523
I. Introduction
II. Lotic Environments
A. The Physical – Chemical Milieu
B. Ecosystem Changes Along the
Stream Course
III. Underground Aquatic Habitats
A. Hyporheic and Phreatic Zones
B. Aquatic Habitats within Caves and
Other Karst Topography
IV. Lentic Ecosystems
A. Geomorphology and Abiotic
Zonation of Lakes
B. Biotic Zonation of Lakes
C. Wetlands, Ephemeral Ponds,
and Swamps
D. Hypersaline Lakes
Literature Cited
I. INTRODUCTION
II. LOTIC ENVIRONMENTS
The contribution of inland waters to the total
biospheric water content is insignificant in terms of
percentage ( 1% according to Wetzel, 2001) but absolutely crucial from the perspective of terrestrial and
freshwater life. Although inland lakes contain 100
times as much water as surface rivers, most lake water
is held within massive basins, such as the Laurentian
Great Lakes of North America, Lake Baikal of Siberia,
and Lake Tanganyika of East Africa. Because most
freshwater invertebrates are clustered within shallow,
well-lighted zones of lakes, the relative importance of
small ponds, creeks, and rivers as habitats for aquatic
invertebrates is much greater than their volume percentages would otherwise indicate. Composition,
species richness, and total density of invertebrates vary
considerably among inland water habitats, as discussed
in this chapter.
Distributions of invertebrates are influenced by interactions among physical, chemical, and biological
characteristics. The general importance of these abiotic
and biotic factors is examined in this chapter. Detailed
information on the ecology of individual taxa in inland
water ecosystems can be gleaned from Chapters 3 – 23.
Flowing waters, or lotic environments, were a principal pathway for evolutionary movement of animals
from the sea to lakes and land. Even today, many taxa
of freshwater invertebrates are restricted to headwater
streams and rivers by unique environmental characteristics of these ecosystems. Compared to nonflowing
waters, or lentic ecosystems, streams are generally
more turbulent than lakes and, therefore, stratification
of the water mass with a thermocline is rare. High turbulence generally maintains high oxygen concentrations, reduces within-stream temperature differences,
and more evenly distributes plankton and suspended
or dissolved nutrients. Temperatures in streams fluctuate over a smaller range than is typical of shallow
littoral zones of lentic ecosystems, where most lakedwelling animals reside (Hynes, 1970). Except in the
northernmost rivers, ice is less commonly encountered
and is generally not as thick as in lakes. Flowing water
habitats frequently possess more habitat heterogeneity,
and the food web in forested drainage basins is more
dependent on allochthonous production (externally
produced plant matter), even though instream production can be important (Thorp and Delong, 1994). Lotic
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
19
20
J. H. Thorp and A. P. Covich
ecosystems are also more permanent on both evolutionary and ecological time frames than most lentic
habitats. Both heterogeneity and permanence are
thought to increase diversity within these ecosystems.
In this section, we review features of epigean, or
surface waters. Characteristics of aquatic environments
where water flows underground (hyporheic, phreatic,
and cave environments) are discussed in Section III.
A. The Physical – Chemical Milieu
1. Basin Morphometry and Characteristics
of Flowing Water
Although stream invertebrates are influenced directly
by the physical features of the local habitat and only indirectly by geological forces shaping the morphometry of
the entire ecosystem, ecologists and invertebrate zoologists must consider all those factors when attempting to
understand what controls community composition. Some
elements of stream morphometry that influence aquatic
communities are stream order, channel patterns (e.g., meandering and braiding), erosion, deposition, and formation and characteristics of pools and riffles.
Classifying streams by their order, or pattern of tributary connections, is a simple technique that aids analysis of longitudinal (i.e., upstream – downstream) changes
in stream characteristics within a single catchment; however, it is less useful, and indeed often misleading for
comparing streams in different catchments. For example, communities within streams in the eastern coastal
plain bear no close resemblance to communities within
similar-order streams in the southwestern deserts (when
flooded) (cf. Figs. 1C and 1D respectively). According to
this scheme, as refined by Strahler (1952) and illustrated
in Figure 2, a continuously running, headwater creek
with no permanent tributaries is classified as a first-order
stream (permanency is defined here as not being
ephemeral on a seasonal or other regular basis). Two
first-order streams combine to form a second-order segment; two second-order creeks join as a third-order
stream, etc. Stream order increases only when two
streams of equivalent rank come together; hence, a
third-order stream that flows into a fourth-order system
has no effect on the subsequent numbering scale of the
larger stream. The Mississippi River, the largest river in
North America, attains a stream order of 11 – 13 in its
lower reaches (depending on which maps are used as the
basis for analysis). Unfortunately, this scheme is not as
useful for many streams in arid portions of the continent
where a dry stream bed (Fig. 1D) may be replaced by a
raging torrent in a matter of minutes or a few hours.
Viewed from the air, streams clearly do not flow in
a simple, straight direction for significant distances
(Fig. 3A), nor do stream beds maintain constant depths
for long stretches. Instead, the channel traces a meandering pattern of gentle and sharp bends through the
basin with alternating bars, pools, and riffles (Fig. 4) as
the water travels downstream along a roughly helical
path. The distance between meanders is a function of
several factors, especially stream width. The river channel also has a tendency to undergo braiding (i.e., the division of the channel into a network of branches) in response to the presence of erodible banks, sediment
transport, and large, rapid, and frequent variations in
stream discharge (Gordon et al., 1992).
Water flows in a somewhat helical manner along the
surface toward an undercutting bank where the deepest
pool is usually located and then passes along the bottom
toward a “point” or “alternate” sediment bar on the opposite bank from the pool. Sediment is continually being
eroded from some areas (e.g., undercut banks and the
head of islands) and deposited in other regions (such as
on bars and the foot of islands). Successive deepwater
pools are separated by shallow riffle regions near the
midpoint of two pools. This sequence of bars, pools,
and riffles is characteristic of both straight and meandering segments of most rivers, especially those with poorly
sorted loads. (See Calow and Petts, 1992; Gordon et al.,
1992; and references cited therein for additional details
on fluvial geomorphology).
Scientists have appreciated the influence of alternating pools, riffles, and (to a much lesser extent) sediment bars for some time. In comparison to pools, riffles
are characterized by greater habitat heterogeneity, sediment size, stream velocity, and sometimes oxygen content. In general, well-flushed, stony riffles support more
species and individuals than silty reaches and pools
(Hynes, 1970). This richness is a consequence, in part,
of the different adaptations required to live in these
physically distinct areas, especially with regard to dissolved oxygen.
Our initial separation of aquatic ecosystems into
flowing- and standing-water environments recognizes
the fundamental importance of this character to the biotic composition of inland water environments. Scientists measure water movement in streams in several
ways, including stream discharge, turbulence, and velocity. Mean and peak values of stream discharge influence various ecosystem characteristics such as substrate
particle size, bed movement, nutrient spiraling, and
rate of animal drift. Stream discharge is calculated by
the equation
Q wdv
where “Q” is the discharge in cubic meters per second
(cms) or cubic feet per second (cfs), “w” the width, “d”
2. An Overview of Freshwater Habitats
A
B
FIGURE 1 Lower-order streams: (A) Rocky-bed mountain stream in Glacier National Park, Montana;
(B) rocky-bed stream from the Smoky Mountains of North Carolina; (C) sandy, coastal plain stream
from South Carolina; (D) intermittent, dry-bed stream from Arizona (photographs by J. H. Thorp).
21
22
J. H. Thorp and A. P. Covich
C
D
FIGURE 1
(Continued )
the depth, and “v” the velocity. As stream flow increases, shear stress is magnified. When the stress
reaches a critical point, the substrate is picked up,
rolled, or generally moved downstream. Various aspects of stream discharge, including mean annual discharge and yearly and historical extremes, are described annually for most rivers in the United States in
the Water Data Reports published by the U.S.
Geological Survey. This information is derived from
multiple gaging stations located near the mouth of each
river and often extending into the headwaters; waterquality data are also available for a few gaging stations
in some of these rivers. These reports are available in
the government documents section of most major
libraries and are accessible through the USGS Web site.
For more details on hydrologic processes, consult
2. An Overview of Freshwater Habitats
FIGURE 2 Bifurcation of streams illustrating the system for
classiifying lotic ecosystems according to stream order. The maximum
order of the hypothetical stream shown here is sixth.
articles or books on this subject, such as Allan (1995),
Calow and Petts (1992, 1994), Gordon et al. (1992),
Newbury (1984), and Statzner et al. (1988).
In addition to being affected by the mean and
range of stream discharge, freshwater invertebrates are
strongly influenced by water velocity. An invertebrate’s
ability to maintain its position on a rock and the
amount of food passing its way (especially for suspension feeders) are both affected by current velocity. The
mean current velocity of a stream at any given time
usually occurs at a depth of about 0.6 of the maximum
depth, as measured downward from the water surface
(or 0.4 of the maximum depth as measured upward
from the stream bottom). The current velocity diminishes progressively toward the bottom from that point
and drops dramatically near the substrate in a 1- to 3mm-thick area known as the boundary layer. For this
reason, velocities experienced by two invertebrates living within boundary layers of separate streams — one
with high and the other with low average current
velocity — may not differ appreciably. Many stream
invertebrates are partially protected from dislodgment
by having a low profile that keeps their bodies mostly
within the boundary layer. Other possible adaptations
for life in fast flow include streamlining, reduction of
projecting structures, development of suckers, friction
pads, hooks or grapples, and secretions of adhesive
products.
Under normal flow conditions, the average current
velocity varies from the headwaters to the mouth of a
23
river. Although many people believe that headwater
streams flow faster than large-order rivers, this popular
idea has been repeatedly disproved (e.g., Leopold, 1953;
Ledger, 1981). This erroneous view of streams may have
arisen because the waters of a turbulent headwater
stream (Figs. 1A and 1B) can “appear” to casual observers to be moving faster than the waters of a higher
order, laminar flow river (Fig. 3A), unless the large river
is shallow and thus more turbulent (Fig. 3B). It is important to note, however, that flow within microhabitats in
any river order can substantially exceed the average current velocity of the stream reach.
Fluctuating current velocity rather than constancy is
typical of all lotic systems. Except in the largest rivers,
zero or near-zero flow rates have been recorded at least
once in most lotic environments since the U.S. Geological Survey began recording stream discharges in the 19th
century. The smaller the stream and the drier the surrounding area (e.g., desert streams), the more likely that
a lotic system will be intermittent on at least a seasonal
basis. At the other extreme, all streams have peak discharges on seasonal and multiyear bases. Flood events,
or spates, may be very predictable, such as those produced by seasonal snow melt in mountainous streams,
or they may occur dramatically over unpredictable time
periods (Resh et al., 1988). Current velocities during
floods and normal periods are related to the width,
depth, and roughness of the stream bed, but rarely
exceed 3 m/s even during floods (falling water seldom
surpasses 6 m/s) (Hynes, 1970). Above 2 m/s, most
rivers begin to enlarge their beds by erosion, unless constrained by rock banks or human construction.
Invertebrates are often adapted morphologically or
in life-history traits to moderate, seasonally predictable
spates, but large unpredictable floods may eliminate all
or most of the fauna of a stream by causing bed movement and severely scouring the bottom. When these
scouring events occur, the sources for recolonization
may be assemblages found in the adjacent hyporheic
zone, in surface waters located upstream or downstream, or in separate, nearby stream catchments (Gibert
et al., 1994; Jones and Mulholland, 2000).
2. The Importance of Substrate Type
The nature and provision of substrates are of
prime importance to survival because the overwhelming majority of lotic invertebrates are benthic (see references in Hynes, 1970; Allan 1995). Substrates provide
sites for resting, food acquisition, reproduction, and
development (e.g., places for pupal case attachment), as
well as refuges from predators and inhospitable physical conditions. Hard substrates are generally valuable
only for their surface qualities (except for species that
bore into wood), whereas relatively soft substrates
24
J. H. Thorp and A. P. Covich
A
B
FIGURE 3
(A) Ohio River above Louisville, Kentucky; here the river is meandering with a
sandy-gravel bottom and a more laminar flow; (B) Rio Grande near Santa Fe, New Mexico; here this
large river is more turbulent than the Ohio River (photographs by J. H. Thorp).
(ranging from sand to silt) provide a three-dimensional
environment. The desirability of a given substrate type
can be evaluated for a specific taxon using a combination of at least the following interrelated characteristics
(not necessarily in order of importance):
1. Whether it is mineral (ranging from large
boulders to fine clay particles), living (algae,
mosses, vascular plant stems and roots, and,
rarely, animal hosts), or formerly living
(e.g., abscised leaves and snags from tree limbs
and trunks).
2. The silt and organic content of, or on, the
substrate, including whether it is bare or
covered by significant amounts of algae, mosses,
or vascular plants.
3. Its particle size, surface texture, and porosity.
4. Its stability, as judged by its retention in the
2. An Overview of Freshwater Habitats
FIGURE 4
Schematic drawing of a stream showing straight reaches
and meanders. In both sections of a river, a sequence of deep (pools)
and shallow (riffle) areas is manifested. Point bars and alternate bars
develop as bed material is deposited in areas opposite from the pools.
The thalweg, a line connecting the deepest part of the channel,
migrates back and forth across the river.
system (e.g., leaves are a short-lived substrate,
whereas boulders are retained almost indefinitely) and its tendency to shift position (ranging from almost constantly shifting sand to
boulders that rarely turn).
5. The physical-chemical milieu in which the
substrate resides, such as the water depth at
which it is located, the ambient oxygen
content, and the range of current velocities
surrounding it.
Even if a particular site meets all the physical and
chemical criteria for a desirable substrate, it may not be
appropriate for an individual invertebrate if a competitor has preempted the space or if it is located in an area
that would make a resident especially susceptible to
predators or abiotic disturbances. For these reasons, the
distribution of a species across a spectrum of substrate
types may not reflect so much the selectivity of the invertebrate species as the effects of substrate limitation.
While a particular taxon may not benefit greatly from
substrate heterogeneity, the species diversity of the entire
community tends to rise with an increase in habitat complexity. In a like manner, an individual may benefit from
maximum substrate, but the community as a whole may
flourish better with some disturbances. This latter concept, embodied in part by the “intermediate disturbance
hypothesis” (Connell, 1978; Ward and Stanford, 1983;
Thorp and Cothran, 1984), suggests that intermediate
levels of biotic or abiotic disturbances (e.g., frequency of
substrate shifting) can promote maximum species diversity under certain circumstances. For a more detailed
discussion of the general effects of substrate characteristics, the reader should initially consult Allan (1995).
3. Thermal and Chemical Constraints on Distribution
a. The Thermal Environment Because temperatures
vary on a seasonal basis and a gradient exists from
headwaters to mouth, it is not surprising that the distri-
25
bution and abundance of freshwater invertebrates are
correlated with the annual mean and range of temperatures (e.g., Hynes, 1970; Ward, 1986). A general stream
temperature pattern on an annual basis is a progressive
increase from headwaters to mouth, especially in
streams originating in the mountains and fed primarily
by terrestrial runoff. [This general pattern may not hold
for low-gradient, north-flowing streams.] Streams originating from coldwater springs, however, with their relatively constant temperatures in the headwaters, can be
warmer in winter near their source than somewhat farther downstream; in the summer, the opposite pattern
exists (Minckley, 1963). In many temperate zone rivers,
annual fluctuation in temperature increases downstream, while diel temperature changes peaks in mid-order reaches (Statzner and Higler, 1985). In contrast, diel
and annual temperature amplitudes are very low in
middle and lower reaches of tropical streams, although
high-elevation, tropical streams have significant seasonal temperature gradients (Covich, 1988).
Predictions of thermal patterns must be modified
to account for effects of tributaries (often colder) and
dams on downstream reaches. Likely differences in
thermal regimes between natural and regulated rivers
are described by the “serial discontinuity concept”
(Stanford et al., 1988), which predicts that the thermal
character of the stream below a dam will be “reset” toward that typical of reaches above the dam. This
pattern is evident if you calculate the number of
degree-days (sum of the average daily temperatures
over a period) in stream reaches above and below the
dam. The timing of insect emergence, an invertebrate’s
tolerance of specific habitats, and annual productivity
of individual species and communities are all influenced
directly or indirectly by the number of degree-days in
those habitats (Sweeney and Vannote, 1978).
Species richness and the nature of invertebrate fauna
vary along a continuum from headwaters to the mouth.
Crustaceans (benthic and planktonic), molluscs, and fish
increase in abundance and diversity with stream order,
whereas insects generally decrease downstream. Because
of previously noted patterns for temperature means and
fluctuations with stream order, it is not surprising that
aquatic ecologists saw a link between thermal regimes
and species diversity. Temperature may influence organisms directly or have an indirect effect through changes
in oxygen saturation levels (the oxygen-carrying capacity
of water decreases at higher temperatures). While it is
true that a great many lotic invertebrates are more
stenothermal (i.e., surviving only in a narrow temperature range) than their lentic counterparts (Hynes, 1970),
it has not been demonstrated that the relationship between invertebrate diversity and temperature is more
than coincidental. The problem in distinguishing
26
J. H. Thorp and A. P. Covich
causative factors is that many biotic and abiotic parameters other than temperature also change along river gradients. For example, habitat complexity, which has often
been demonstrated to significantly influence diversity,
varies downstream, often reaching a maximum value in
mid-order sections of rivers.
b. The Chemical Environment Certain natural chemical features of streams and lakes significantly affect
invertebrate abundance and diversity by influencing
habitat quality. Of these, the most crucial are probably
dissolved oxygen and salinity (water hardness). Hydrogen ion concentration, or pH, is occasionally important
in natural systems. Anthropogenic pollution has had a
severe impact on the integrity of aquatic ecosystems by
affecting these chemical parameters as well as by introducing organic and inorganic toxicants.
As a general rule, dissolved oxygen decreases
downstream. This pattern occurs because the upper
reaches of a river are usually more turbulent, the surface to volume ratio is more favorable for diffusion,
and temperatures are lower. In the last case, oxygen
solubility is related in an inverse, nonlinear manner to
temperature; this gas is markedly more abundant in
cold waters. Pure water in equilibrium with air at standard pressure holds 12.770, 10.084, 8.263, and 6.949
mg O2/L at 5, 15, 25, and 35°C, respectively (see Mortimer, 1981, in Wetzel, 1983, p. 158). Because oxygen
is produced and respired by plants, it may be locally
more abundant or even supersaturated during the day
in regions with heavy plant growth. However, oxygen
potentially decreases at night to lethal levels in weedchoked streams because of high oxygen demands for
decomposition and plant respiration. The probability
of lethal oxygen concentrations occurring for the invertebrate species typical of a stream reach rises as currents diminish; hence, the problem is more serious in
lakes and seasonally sluggish streams than in fast-flowing lotic habitats.
Invertebrates differ both in their respiratory needs
and in their anatomical and behavioral adaptations for
obtaining oxygen. Respiratory rates of lotic species are
generally dependent on external oxygen concentration
(tension) at medium to low levels, while oxygen consumption rates of animals in lentic or typically sluggish
lotic environments tend to be independent of ambient
tensions, at least at medium-to-high concentrations
(Hynes, 1970). Currents provide a steady renewal of
oxygen to the respiratory surfaces of invertebrates.
Where velocities are too low, the animal may need
to ventilate its gills or obtain atmospheric oxygen. For
example, crayfish normally move water over their
respiratory surfaces with “gill balers” but some species
climb partially or completely out of water if oxygen
tensions drop too low (returning when gills need moistening). Because invertebrates have different oxygen
requirements, it is tempting to assume that interspecific
differences in distribution are partially caused by oxygen
limitations (e.g., contrasting assemblages of caddisflies
in mountain streams and floodplain rivers). Certainly
many stream animals, such as larvae of the blackfly
Simulium, cannot obtain adequate oxygen in still water
(Hynes, 1970), but this explanation for restriction from
lentic habitats is less effective in resolving species differences in distribution along the entire stream reach. Even
if one demonstrated in the laboratory that the oxygen
tensions in high-order rivers were inadequate to support
a caddisfly from a mountain stream, this is not evidence
that oxygen was the principal factor leading to the present pattern of distribution. Other factors, such as interspecific competition, resource type (e.g., seston particle
size), substrate form, or predators, may have been initially responsible, with fine tuning of respiratory adaptations coming later.
The ionic content of inland waters is usually extremely dilute in comparison to seawater, with four
cations (Ca2, Mg2, Na, and K) and four anions
2
2
(HCO
3 , CO3 , SO4 , and Cl ) representing the total
ionic salinity in most freshwater systems, for all practical purposes (Wetzel, 2001). Soft waters are generally
found in catchments with acidic igneous rocks, whereas
hard waters (relatively high salinities for freshwater)
occur in drainage regions with alkaline earths, usually
derived from calcareous rocks. The global mean salinity of river water is 120 mg/L; North American waters
are slightly harder, averaging 142 mg/L [from Livingstone (1963) and Benoit (1969) in Wetzel (1983, p.
180)]. Most soft waters in North America have salinities 50 mg/L. Salinities are slightly elevated during
periods of low flow, and they generally increase downstream because of bedrock erosion (Hynes, 1970) and
human influences on watersheds.
Except for a few hypersaline environments, such as
mineral springs and alkaline and brine lakes, inland waters are hypotonic to the internal fluids of freshwater
species; therefore, all freshwater animals need some
ability either to osmoregulate (almost all species) or, at
the minimum, to control the relative abundances of certain ions (necessary even for freshwater Cnidaria; see
Chapter 5). Because animals with either partially calcareous exoskeletons (e.g., crayfish, ostracodes, and
cladocera) or hard calcareous shells (snails and mussels)
have an added need for calcium ions, one might expect
them to be particularly excluded from soft-water habitats (see Chapter 18). Indeed, molluscs are rare in softwater lakes and less abundant in soft-water streams, but
this limitation is not as severe as in lentic ecosystems.
Molluscs can live in some soft-water streams, because
2. An Overview of Freshwater Habitats
they can tolerate extremely dilute tissue fluids (the lowest of any metazoa; see Chapter 11). The minimum concentration of Ca2 tolerated by mussels (2 – 2.5 mg
Ca/L) is just adequate to prevent the rate of shell dissolution from exceeding the rate of shell deposition. Despite their heightened need for calcium, crayfish can be
abundant in many soft-water streams of the Southeast
where these ions are difficult to obtain. The problem of
ionic loss at the critical period when the old exoskeleton
is molted is partially overcome by the crayfish habit of
consuming its shed exuvia as a way of extracting additional calcium ions. This behavior and the efficient osmoregulatory system of the crayfish (with gastrolith formation) allow this decapod to colonize a greater
chemical range of habitats (see Chapter 23).
In the last decade, the catastrophic effects of acid
precipitation and mine drainage on freshwater ecosystems have drawn great attention from scientists and
politicians alike, but relatively unpolluted rainwater
and runoff from thick forest litter are also slightly
acidic (though not at lethal levels). Bogs and the
streams draining them are among the most naturally
acidic ecosystems; sphagnum bogs usually have a pH
below 4.5 and occasionally below 3.0 (Wetzel, 2001).
Streams draining limestone catchments are well
buffered by carbonates and therefore have moderate to
high pH values as well as plentiful supplies of calcium.
In contrast, acidic waters are normally low in calcium,
causing potential osmotic problems for their residents.
Despite probable ionic and pH difficulties which may
have eliminated certain taxa, naturally acidic ecosystems often support a surprisingly diverse invertebrate
assemblage.
B. Ecosystem Changes along the Stream Course
1. Patterns along the Continuum
Physical and biological characteristics clearly
change from headwaters to the river mouths. The river
widens, deepens, and becomes more turbid downstream, often removing part of the bottom from the
photic zone. Stream shading from riparian trees lessens,
with a corresponding increase in percentage of the surface receiving direct solar radiation. Concurrently, the
relative importance values for autochthonous (internally produced) and allochthonous carbon to energy
budgets alter along with the ratio of production to respiration. As a result of these and other habitat changes,
the invertebrate fauna undergoes dramatic shifts in
species composition, relative abundances, and functional feeding groups.
This holistic view of the river was not synthesized
until formation of the river continuum concept (RCC)
by a group of aquatic ecologists working in temperate
27
deciduous streams of North America (Vannote et al.,
1980; Sedell et al., 1989). This important theory
has proved to be one of the most hotly debated in
recent decades, and that fact alone contributes to
the pivotal role it has played in aquatic ecology. Individual hypotheses in the RCC have received both support
and criticism (as examples of the latter, see Winterbourn, 1982; Statzner and Higler, 1985; Junk et al.,
1989; Thorp and Deong, 1994). It is not our purpose to
defend or assail this holistic, ecosystem concept; instead, we wish to acquaint you with some of its predictions as they could influence your understanding of the
habitat and ecology of lotic invertebrates. Because the
RCC was originally developed from studies of relatively
undisturbed, North American streams with forested
headwaters (cf. Fig. 1B), the following comments may
not be as applicable to stream invertebrate communities
in other biomes or regions of the world.
The river continuum concept relies on three theoretical pillars of support. The first principle is that
stream communities originate in response to a continuous gradient of physical variables present from
headwaters to the mouth; this view contrasts with a
previous portrayal of natural streams as a series of
disjunct communities proceeding downstream in a stepwise fashion. The second canon is that biotic communities within the “wetted channel” of the river cannot be
divorced from either the adjacent riparian zone or the
surrounding biotic and geomorphic catchment area
that funnels water, nutrients, and other material into
the aquatic ecosystem. The final tenet is that the nature
of a downstream assemblage is inextricably linked with
processes occurring upstream (this has been challenged
for floodplain rivers by Junk et al., 1989).
Of the many physical, chemical, and biological
predictions made by the RCC, the one most relevant to
understanding invertebrate ecology concerns longitudinal changes in the relative abundance of functional
feeding groups and their food resources. The proposed
relationships between stream size and progressive alterations in community composition are illustrated in
Figure 6. As heterotrophs, invertebrates consume living
and/or dead organic matter, processing it in ways characteristic of several “functional feeding groups.” These
groups include “shredders,” which break up coarse
particulate organic matter (CPOM) such as abscised
leaves; “collectors,” which gather or filter CPOM, fine
particulate organic matter (FPOM), and / or live, drifting organisms; “grazers” or “scrapers,” which remove
algae or other aufwuchs growing on rocks or other
substrates; and “predators,” which may include carnivores and some herbivores.
The traditional means of assigning a functional
feeding label to a life stage of a particular species has
28
J. H. Thorp and A. P. Covich
been either to observe the organism’s feeding behavior
(preferably in the field) or to analyze its gut contents on
a seasonal basis (e.g., Benke and Wallace, 1997). This
task is time consuming and may be impractical for
some taxa. References to the food resources and feeding behavior of species or larger taxonomic categories
are reported in many research papers (for examples, see
chapters in this book and the tables and references on
aquatic insects in Merritt and Cummins, 1996). The
danger in relying on general sources, however, is the
well-established observation that functional feeding
classifications can vary significantly with life-history
stage, season, and other ecological features; it is also
difficult to assign omnivores to any single or consistent
functional feeding group (see summary of some limitations of this method in Mihuc, 1997). An approach
that overcomes some of these problems is to construct
food webs using data on stable isotope and/or fatty
acid content of producers (dissolved organic matter
through living terrestrial and aquatic plants) and
aquatic consumers; these data reflect the cumulative
feeding patterns of freshwater invertebrates over periods of up to 3 months (e.g., Thorp et al., 1998).
The RCC predicts that the invertebrate assemblages
of headwater streams, with their heavy canopy of riparian trees, will be dominated by shredders and collectors.
As the stream widens and more light reaches the water
surface, autochthonous production from benthic algae
and rooted macrophytes increases, allowing the development of a significant grazer fauna; shredders should
diminish dramatically in these mid-order reaches of the
river. Farther downstream, the river deepens so that
much of the bottom is below the photic zone (supposedly limiting autotrophic production in the river), and
the channel is hypothesized to be sufficiently wide to
preclude any significant input of allochthonous production from the riparian zone. In this high-order section of
the river, collector – filterers are expected to dominate
the ecosystem. These conclusions on primary sources of
organic matter have been strongly challenged for large
rivers by both proponents of the flood pulse concept
(Junk et al., 1989), who emphasize the importance of
allochthonous production generated on river floodplains, and proponents of the riverine productivity
model (Thorp and Delong, 1994), who stress the
greater importance of instream (autochthonous) primary production to secondary (animal) production, especially at higher trophic levels.
2. Nature of the River’s Source
One weakness in the original formulation of the
RCC was its reliance on data collected from a narrow
range of stream sizes (mostly stream orders 1 – 3). Even
today, our knowledge of assemblages living at opposite
ends of the continuum — springs and intermittent
streams versus large rivers — is woefully inadequate
(Meyer, 1990).
Research on intermittent, or ephemeral, streams
(Fig. 1D) is more common in arid regions of North
America (e.g., Matthews, 1988; Forrester et al., 1999),
but has expanded into headwater streams in more humid regions of eastern North America (e.g., Delucchi
and Peckarsky, 1989). Almost all streams are subject to
seasonal and / or aperiodic disturbances from unusually
high current velocities and stream discharges, but only
intermittent streams fluctuate regularly from dry to
wetted basin. When a stream stops flowing, a resident
invertebrate may have several options for its survival or
that of its future progeny: it may move downstream,
seek shelter in deep pools, enter the hyporheic zone
(i.e., the wetted substrate below the surface waters, as
described in Section III.A), emerge as an adult (some
aquatic insects), develop a resistant stage, or lay eggs
and then die. Although the composition of the invertebrate fauna in permanent and intermittent streams
may differ, life-history patterns of invertebrates in
ephemeral streams are not particularly unique (Delucchi and Peckarsky, 1989), perhaps because options for
migrating downstream or entering the hyporheic zone
are readily available for many species. The ecology of
that portion of the ecosystem, however, may be significantly different from that of a community functioning
within permanent streams (e.g., Forester et al., 1999).
There is a tendency to think of streams as originating exclusively from terrestrial runoff, but many headwater systems are derived from springs. Near their
source, spring-fed streams often vary in physical, chemical, and biological features from other types of first-order streams (e.g., Covich, 1988). Temperatures within
spring-fed streams are relatively more constant, and
oxygen concentrations are occasionally lower, sometimes to near-lethal levels for metazoa. In other cases,
hot springs may provide habitats that range into the
upper lethal limits for eukaryotes (see Chapters 19 and
20) and even prokaryotes. Metazoan life near the
upwelling region is rarely possible; but, as ambient
temperatures approach 40°C or lower in side channels
and pools and in areas farther downstream, conditions
become suitable for some invertebrates (e.g., Barnby
and Resh, 1988). These include a very restricted number of species of nematodes, oligochaetes, mites, dipterans (such as brine flies), and ostracodes and a few other
crustaceans. Geothermal springs are often extremely
vigorous environments not only because temperatures
may approach the boiling point but also because their
waters are frequently laden with high concentrations of
sulfur. The ionic content of springs can differ from that
of from nearby streams supplied by runoff from land.
Springs in limestone regions are well buffered and are
2. An Overview of Freshwater Habitats
frequently highly charged with calcium bicarbonate
(Hynes, 1970). Saline springs exist where the ionic content is too high for many invertebrates.
The biota of springs often vary from other firstorder streams as a result of their connection to underground rivers and their aquatic constancy (Gibert et al.,
1994; Jones and Mulholland, 2000). A net placed near
the spring source can capture unusual subsurface
species that have been washed out of their subterranean
environments. In this way, blind troglobitic shrimp have
been retrieved from surface waters in the springs of the
San Marcos River of Texas (Longley, 1986; Strenth
et al., 1988). Because springs are relatively uniform
environments over long ecological periods, it is not unusual to find relict species in these environments that
have survived the retreat of the glaciers only in these
limited habitats (Hynes, 1970). For discussion on endemic groundwater fauna in unglaciated eastern North
America, see Strayer et al. (1995).
3. Large Rivers
At the opposite extreme of the continuum, the invertebrate biota and ecological processes in large rivers
(e.g., Figs. 3A, 3B, and 5) have rarely been studied in
the past (although this is beginning to change) despite
their importance to humans. Hynes (1989) pointed out
that only 4% of all ecological publications of flowing
waters have involved large rivers. Consequently, certain
misconceptions about the nature of large rivers have
arisen, such as descriptions of the “typical” large river
as a slow, meandering, soft-bottomed stream. In con-
29
trast, observations by one of us (JHT) in the Mississippi, Ohio, and Tennessee Rivers suggest that the bottom is more likely to be sand, gravel, or cobble except
in the shallow, slow-moving water near banks and in
the mouths of tributaries where silt often accumulates.
Furthermore, evidence for differences in meandering
patterns along a river continuum has not yet been generated, and in comparison to upstream reaches, the current velocity is generally higher in large rivers. Because
governments have been removing fallen trees and regulating both flow and channel depth in navigable rivers
since the mid-1800s, the present nature of these ecosystems may be considerably different from the original
state (Minshall, 1988). For example, removal of snags
has greatly reduced the abundance of shallow-water
hard substrates, thereby influencing composition of
both the insect assemblage and the fish that normally
fed upon these insects (cf. Benke et al. 1994).
Many aquatic ecologists lacking experience in large
rivers (other than reservoirs) often think of them as
“lakelike,” perhaps expecting similar physical and
chemical conditions. Unlike lentic environments, however, the helical flow of riverine currents prevents formation of thermoclines and major zones of oxygen depletion within the water column; this has considerable
influence on processes of nutrient cycling (or spiraling
in rivers). The greater depth and turbidity of sedimentladen rivers reduce the amount of benthos receiving
light so that only a narrow photic zone exists. The
resulting restriction on phytoplankton net production,
along with microbial breakdown of allochthonous
FIGURE 5 Photograph of a large river (the Ohio River upstream of Louisville, Kentucky),
showing the laminar flow nature of its waters (photograph by J. H. Thorp).
30
J. H. Thorp and A. P. Covich
detritus, is thought to be a primary factor producing a
heterotrophic metabolic state in most large rivers
(Howarth et al., 1996). The potential importance of
physical factors in an advective environment (Pace
et al., 1992) and their influence on the role of biotic
interactions (Thorp and Casper, unpublished data) are
currently being investigated.
The RCC appears generally correct in predicting
the important role for suspension feeders in large
rivers, but it underestimates the role of some other
groups, such as grazers. As predicted by the RCC, suspension feeders predominate in both the benthos (e.g.,
mussels and midges) and pelagic zones (various crustacean and rotifer zooplankton) of large rivers, especially since the invasion of zebra and quagga mussels
into eastern rivers (Thorp et al., 1998). However, the
predicted absence of grazers, as would be anticipated in
the Ohio River from Figure 6, is certainly incorrect.
Abundant populations of snails in the Ohio consume
various types of benthic algae, other aufwuchs, and detritus (depending on the depth distribution of snails;
K. Greenwood and J. H. Thorp, unpublished data). Invertebrates in general tend to be clustered in nearshore
(especially on rocks, as in Fig. 5), shallow-water regions of rivers (Thorp and Delong, 1994), where they
have access to both benthic algae and allochthonous
material entering from the riparian zone.
Because the North American Benthological Society
focuses on nonpelagic ecology and members of the
American Society of Limnology and Oceanography are
primarily oriented to lentic and oceanic environments,
studies of plankton ecology in rivers are relatively rare
in North America, though more common in Europe.
However, the increasing importance of zebra mussels in
rivers has contributed to studies of these potamoplankton (from the Greek potamos, or river) and
benthic – pelagic coupling processes (e.g., Pace et al.,
1998; Jack and Thorp, 2000; Thorp and Casper, unpublished data).
One of the more interesting trends in large-river research, that seems to be developing includes freshwater
zooplankton. This trend revolves around the importance of lateral slack-water areas, or storage zones
(Reckendorfer et al., 1999, Thorp and Casper, unpublished data), to community and ecosystem processes in
the river as a whole.
III. UNDERGROUND AQUATIC HABITATS
A. Hyporheic and Phreatic Zones
FIGURE 6 Predicted relationship between stream size and
structural and functional attributes of a freshwater ecosystem as
embodied in the river continuum concept. P/R refers to the ratio of
production to respiration (from Fig. 1 in Vannote et al., 1980).
Until recently, aquatic ecologists considered the
boundaries of rivers to extend only from the air-water
interface down to the silty, sandy, or rocky “bottom”
of the river. We now realize that this definition is overly
limiting and view streams as four-dimensional ecosystems with longitudinal (upstream-downstream), vertical (deeper into the sediments), lateral (floodplain), and
temporal components (cf. Ward, 1989).
Early this century, scientists recognized the potential importance of interstitial spaces within sediments
of lakes and streams as refuges for small animals. This
interstitial habitat is termed the “hyporheic zone”
within lotic systems and the “psammon zone” in lentic
environments. The depth of the hyporheic zone varies
greatly according to bed topography, substrate porosity, and water velocity (Boulton et al., 1998), being
greater, for example, in rivers having a gravel bed
rather than a mud bottom (the habitable interstitial
space is influenced by, but is not directly related to,
sediment porosity). The overlying surface, or epigean,
waters strongly influence the nutrient content, oxygen
tension, and other physical and chemical characteristics
of this zone and, consequently, the nature of its fauna,
2. An Overview of Freshwater Habitats
or “hyporheos.” Likewise, upwelling from the hyporheic zone can provide nutrients to stream organisms. While many organisms dwell much of their lives
in the hyporheic, it is not clear whether this zone serves
as the refuge from stream spates that it was once
thought to be (Gayraud et al., 2000).
Almost all freshwater invertebrate phyla have representatives in hyporheos, with the probable exceptions
of Cnidaria and Porifera. They include several orders
of insects (e.g., midges, elmid beetles, and stoneflies),
mites, crustaceans (including cladocera, ostracodes,
and amphipods), oligochaetes, turbellarians, rotifers,
snails, tardigrades, and protozoa.
Hyporheic organisms are usually abundant in
streams unless the substrate is bedrock, clay, or other
material containing pore sizes under 50 m (Williams,
1984), but their densities reflect differences in organic
content and oxygen, among other factors. Below a few
centimeters, the density of hyporheos is generally inversely related to substrate depth, with very few individuals occurring much below 1 m into the sediment.
Because the hyporheic food web is based on detrital inputs from surface waters (Boulton et al., 1998), animal
density is strongly related to the amount of organic carbon present in the interstitial spaces (Brunke and
Gonser, 1999). Organisms in the hyporheic, and presumably the phreatic, are more resistant to low oxygen
conditions (hypoxia) than are epigean species, but they
cannot permanently tolerate suboxic conditions
(Malard and Hervant, 1999). The distribution and local diversity of hyporheic invertebrates are also influenced by historical patterns of glaciation (Strayer and
Reid, 1999).
The hyporheic zone also extends laterally into the
stream bank, where often the distance it penetrates
horizontally from the river proper, is similar to that it
penetrates vertically into the river bed. Data from the
Flathead River, a gravelly stream of the northern
Rocky Mountains, have shown, however, that the hyporheos permeates groundwater as far as 2 km from
the surface water of the river (Stanford and Ward,
1988). It has been known for some time that a unique
fauna can occupy the phreatic zone — a saturated underground area where the water moves slowly laterally
in the absence of direct influence from surface water
rivers. The study by Stanford and Ward, however,
demonstrated that the groundwater community within
up to 2 km from the Flathead River was hyporheic
rather than phreatic because there were physical
(flow), nutrient, and biotic connections between the
two areas. The biota of the adjacent phreatic zone was
dominated by subterranean crustaceans, while the hyporheic zone supported stoneflies and other typically
riverine taxa.
31
B. Aquatic Habitats within Caves and Other
Karst Topography
Regions of the world containing large amounts of
soluble limestone and adequate underground water often form complex subterranean cavities; such karst
topography is widespread in North America (Fig. 7).
Karsts develop when rocks are fractured and are especially susceptible to solution (e.g., crystalline, high
calcite limestone). Over time, small cavities can enlarge
to produce huge underground caverns. The public generally becomes aware of solution processes only when
an above-ground entrance exists or a sinkhole develops
(also called sinks or, more properly, dolines).
Caves can be defined as karsts with an entrance,
passageways, rooms, and a terminal blockage to all but
water and material it carries. Cave environments provide habitats (Fig. 8) for many types of invertebrates
which differ in several significant ways from surface, or
epigean, aquatic habitats. They have very constant temperatures, that are equal to the average yearly surface
temperature of the local geographic region, light is dim
or absent, and abiotic disturbances are probably less
frequent and severe (although floods occur in caves).
Food is quite scarce in caves, as it usually arrives
only through groundwater or periodic movement of animals that seek refuge within an open cave after having
fed outside (e.g., bats which deposit organic guano in the
cave). The principal forms of organic matter supporting
the food web are particulate (POM) and dissolved organic matter (DOM) along with heterotrophic bacteria
growing in water or on suspended and benthic detritus
and some chemoautotrophic bacteria. Consequently,
food webs are relatively simple, consisting mostly of detritivores and their predators. The density and species diversity of a single cave community are usually quite low.
In contrast to the simplicity of a cave community,
the species composition of one cave can be markedly
FIGURE 7
Karst topography in the 48 contiguous states of the
United States (from Ritter, 1986).
32
J. H. Thorp and A. P. Covich
FIGURE 8 (A) A cave habitat in southern Indiana; (B) The blind, troglobitic crayfish Orconectes
testii (both photographs courtesy of H. H Hobbs, III).
different from that of an adjacent but unconnected
neighboring cave. Endemism is especially prominent in
caves because of inherent geographic isolation. The
species radiation of amphipods in North America has
been much greater in karsts than in surface waters (see
Chapter 19). A total of 927 species of invertebrates
that live exclusively in caves or associated subterranean
habitats in the contiguous 48 states in the United
States, have been described so far, with more than 50%
of the aquatic species and subspecies derived from only
18 counties representing less than 1% of the land area
(Culver et al., 2000)! Just over 50% of these subterranean species are from only a single county, and 95%
of the cave species are listed by the Nature Conser-
2. An Overview of Freshwater Habitats
vancy as vulnerable or imperiled. It should be evident,
therefore, that habitat protection at those sites is absolutely critical to species survival.
Obligate cave species, or troglobites (as opposed to
the facultative troglophiles), usually have low metabolic rates and a much longer lifespan than their surface relatives. They are also frequently blind and lack
body pigmentation. Reproductive yield is spread over
longer periods with fewer but more yolky eggs formed
during each reproductive bout. For example, a female
crayfish of the troglobitic species Orconectes inermis in
Indiana caves may carry only a few dozen large eggs,
while nearby epigean, congeneric individuals can bear
hundreds of small eggs (J. H. Thorp, personal observation, and H. H. Hobbs, III, personal communication;
see also ecologically similar species in Fig. 8).
Information on karst communities can be gleaned
from journals such as the National Speleological Society Bulletin. See also Chapters 19 and 23 in this book
for details about cave amphipod and decapod crustaceans, respectively. For other information on the
ecology of groundwater communities, see specialty
texts such as Gibert et al. (1994) and Jones and Mulholland (2000).
IV. LENTIC ECOSYSTEMS
Unlike the situation for lotic habitats, the chemistry,
physics, geology, and biology of lentic (or lacustrine)
environments are well covered in several standard limnology textbooks (e.g., Cole, 1994; Lampert and Sommer, 1997; Wetzel, 2001), most of which devote very
little space to stream studies. Lake limnology is also discussed in more depth in various specialty texts (e.g.,
Taub, 1984; Carpenter, 1988) and in four volumes on
limnology by Hutchinson (1957, 1967, 1975, 1993).
For this reason, our discussion of lake habitats is briefer
than that accorded to stream ecosystems.
A. Geomorphology and Abiotic Zonation of Lakes
Although rivers and streams are important biologically, more than 99% of the inland surface waters are
in fresh and saline lakes. These standing water, or
lentic, ecosystems occur on every continent but are
concentrated in the formerly glaciated regions of the
Northern Hemisphere. Except for ~20 lakes with
depths 400 m, most natural and human constructed
lentic systems have average depths of 20 m (Wetzel,
2001). Most are geologically young, dating from the
last glacial period in the case of natural lakes or from
the last century for human-made lakes. According to
Hutchinson (1957), natural lakes are formed by over
33
70 distinct processes. Most result from catastrophic
phenomena (landslides and glacial, tectonic, and volcanic activities), but some form less violently from the
action of rivers (e.g., oxbow lakes), waves, and rock
solution. Before the advent of extensive, commercial
fur trapping in North America, ponds built by beavers
were a pervasive feature of the headwaters of many
North American streams (Naiman et al., 1986).
Lentic ecosystems can be divided into several abiotic zones, primarily based on distance from shore,
light penetration, and temperature change (Fig. 9). The
photic zone extends downward to the depth of 1%
light penetration; all primary producers and most heterotrophic animals live within this zone. Areas below
these depths are variously called the aphotic or profundal zone (the latter often in reference to benthos). The
shallow, nearshore region of the photic zone where
rooted macrophytes exist is termed the littoral zone.
The entire mass of open water located away from both
the shore and the littoral zone is known as the limnetic
or pelagic zone.
On a seasonal basis, most lentic ecosystems in
North America become stratified with a layer of lighter
water, the epilimnion, floating over the denser hypolimnion (Fig. 9). At least during the summer, the
epilimnion is considerably warmer than the hypolimnion. These two zones are separated by a layer of
rapid temperature change called the metalimnion. The
boundary between the epilimnion and the hypolimnion, where temperature changes occur most
rapidly with depth, is defined as the thermocline (it is
typically within the metalimnion). Because there is minimal exchange of water between upper and lower zones
during stratification, the hypolimnion is frequently
lower in oxygen, higher in nutrients, and different in
pH and other chemical concentrations. When lake
stratification breaks down for a short period during
one or more seasons, much or all of the water mass recirculates in a process referred to as lake turnover. If a
lake mixes completely twice a year, it is referred to as
dimictic; if it turns over only once a year it is called
monomictic. In other lentic systems, complete mixing
either rarely takes place (in oligomictic lakes) or occurs
more than twice per year (polymictic lakes). In addition
to thermal zones, some lakes are chemically stratified
(commonly with a heavier layer of saline water in the
hypolimnion). Some of these lakes never mix completely and are termed meromictic; they only circulate
in the upper zones. As a consequence, they are typically
devoid of oxygen in deeper zones, where nutrients can
accumulate over time.
The benthic zone can be further divided into somewhat arbitrary categories based on other ecosystem
characteristics, such as substrate type, wave action, and
34
J. H. Thorp and A. P. Covich
FIGURE 9
Biotic and abiotic zones within a lake along with a list of some representative freshwater
invertebrates found within these zones.
vegetation type. The importance of substrate type for
lentic environments is similar to that described earlier
for lotic ecosystems.
fleas (cladocera; Chapter 20), alternate periods of resting or foraging on the bottom with intervals of swimming and foraging in the water column.
1. Neuston
B. Biotic Zonation of Lakes
Lake invertebrate assemblages can be subdivided
primarily into “zooplankton”, within the water column, and “benthos,” which live in, on, or just above
the bottom (commonly in association with macrophytes). A third, smaller group encompasses the
“neuston,” which live at the air – water interface. These
first two categories remain quite distinct except in three
general cases. A few species, such as the phantom
midge Chaoborus, migrate upward at dusk into the
plankton and return to the benthos near dawn. Meroplankton, in contrast to the much more diverse holoplankton, spend only a portion of their life cycle in the
water column. For example, chironomid midges are
benthic for most of the larval period (the short-lived
adults are aerial), but swim within the water column
during the earliest stages of their larval existence.
Dreissenid mussels, which invaded North America in
the late 1980s, have a veliger stage that is planktonic
for up to 3 weeks (Garton and Haag, 1993; see also
Chapter 11). Although not true members of the plankton, many nearshore species, such as littoral zone water
The term neuston refers to the assemblage of organisms associated with the surface film of lakes,
oceans, and slow-moving portions of streams. It generally includes species that live just underneath the water
surface (hyponeuston), individuals that are above but
immersed in the water (epineuston), and taxa that
travel over the surface on hydrophobic structures (superneuston or, more properly, a form of epineuston).
This name is similar to, or a subset of the older name,
pleuston (sometimes neuston is used in reference to the
microscopic components of the more encompassing
pleuston). The density of neustonic organisms decreases with increasing turbulence.
The neustonic food web is primarily supported by a
thin bacterial film on the upper surface of the water, a
concentration of phytoplankton near the surface, and
allochthonous inputs from trapped terrestrial and
aquatic organisms. Protozoa are common in this assemblage (which also includes algae and floating macrophytes), whereas other typically planktonic taxa are
rare (one exception is the cladoceran Scapholeberis).
Moving over the water surface are springtails (Collem-
2. An Overview of Freshwater Habitats
bola), some arachnids (mites and water spiders), and
various families of heteropteran bugs (e.g., water striders, Gerridae) (refer to Chapter 16 for information on
arachnids and to Chapters 17 and 18 for details about
insects and springtails). Larger neustonic species are especially vulnerable to predation from both aquatic and
terrestrial predators, and all species must be adapted to
the higher ultraviolet radiation present near the water
surface (see Williamson et al., 1999, for recent perspectives on UV and lake plankton).
2. Zooplankton of Pelagic and Littoral Habitats
Zooplankton include all animals in the water column that float, drift, or swim weakly (i.e., they are at
the mercy of currents). Fish, as powerful swimmers, are
the principal members of freshwater “nekton” in littoral and pelagic environments. Some predatory zooplankton are active swimmers and comprise the “nektoplankton”; examples are the large cladoceran
Leptodora kindtii (Chapter 21) and mysid “shrimp”
Mysis relicta (Chapter 19). Zooplankton thrive within
two distinct habitats: open-water epilimnion and the
nearshore littoral zone. The littoral habitat is the more
heterogeneous of the two in terms of spatial and temporal complexity. The seasonal presence of macrophytic plants (algae and vascular plants) and proximity
of planktonic and benthic habitats provide important
aspects of physical complexity to the littoral zone. Additionally, temperatures fluctuate more rapidly in shallow water, and wave turbulence is greater. Oxygen is
rarely limiting nearshore (except, sometimes in eutrophic lakes), but the effects of ice formation can be
more severe. The majority of lentic research has dealt
with the pelagic zone.
Biological interactions in the two planktonic habitats are different in response to those spatial and
temporal differences and to variance in bases of the
two food webs. Pelagic plankton rely on nutrients
regenerated by lake turnover, offshore transport, and
internal recycling; photosynthesis is exclusively by phytoplankton. Littoral plankton derive nutrients from the
same three sources, but they have first access to allochthonous carbon and nutrients. They can also gain
energy and recycled nutrients from the benthos on a
continual basis (not possible for pelagic zooplankton
except during lake turnover). Phytoplankton, periphyton, and macrophytes provide photosynthetic products
to support the invertebrate assemblage of the productive littoral zone.
Predator – prey relationships and zooplankton behaviors vary between pelagic and littoral zones. Openwater copepods, cladocera, mysids, and a few rotifers
migrate vertically on a diel basis. Fewer littoral zoo-
35
plankton migrate, and among those that do, a typical
pattern is for offshore movement at dusk and inshore
migration at dawn. Ecologists believe that an important cause for both horizontal and vertical migration is
predator avoidance. The types of predators in the two
habitats are considerably different. Aquatic insects in
general are of minor importance in the pelagic zone,
but insect predators, such as dragonflies and predaceous beetles, are extremely abundant nearshore. Although predaceous adult and larval fish are usually
more numerous in shallow, vegetated regions, the
greater habitat complexity of the littoral habitat reduces foraging efficiency relative to open-water regions.
Microcrustacean predators are present in both habitats.
3. Littoral and Profundal Benthos
Benthic invertebrates are often divided into three
arbitrary size classes based on the mesh dimensions of
the sieve used to process the samples. Macroinvertebrates are retained by coarse sieves ( 200 m mesh)
and are usually sorted with No. 60 or 35 U.S. Geological Sieves (250 and 500 m mesh, respectively). Meiofauna, which are plentiful but rarely identified, need to
be collected with fine sieves ranging from 40 to 200
m mesh (usually 100 m); microbenthos pass even
these minute openings. The effort required to process
benthic samples, especially from vegetated and / or silty
habitats, increases tremendously with diminishing mesh
size; consequently, species smaller than macrofauna are
often ignored. While these size categories in themselves
have no natural ecological or taxonomic significance,
the frequent exclusion of the smaller forms from ecological analyses may have severely influenced our
perception of how benthic assemblages function. For
example, Strayer (1985) estimated that the micro- and
meiobenthic animals contributed 68% of the species,
98% of the individuals, 25% of the biomass, and 35%
of the production of the zoobenthos in Mirror Lake,
New Hampshire. Studies in Mirror Lake revealed that
over half of the benthic micro- and meiofauna live in
the top centimeter of sediments (Strayer, 1985, 1986).
Most freshwater benthos reach their maximum
densities and diversity in shallow water and decline perceptibly with increasing depth in the profundal zone.
Few macroinvertebrates tolerate conditions in the profundal zone beneath the seasonal thermocline, but micro- and meiofauna can be abundant in deep water.
This pattern probably reflects gradients of oxygen availability, habitat heterogeneity, and food resources — all
of which are greater in the littoral zone. Studies of vegetated and nonvegetated regions of the littoral zone
demonstrate the great value of macrophytes in reducing
predation rates on benthic macrofauna (e.g., Hershey,
1985). This refuge is especially crucial because benthic
36
J. H. Thorp and A. P. Covich
animals are generally poor swimmers and have difficulty escaping highly motile predators.
A variety of distinctive habitats are available for
benthic invertebrates. Nematodes, gastrotrichs, flatworms, and other meiobenthos frequent infaunal habitats where they move between sediment particles
(Chapters 6 – 9). Those living on or among sand grains
are called psammon or interstitial organisms.
Macroinvertebrates, such as freshwater mussels,
oligochaetes, and some crayfish, regularly burrow in
the substrate. Many motile and sedentary species live
on the surface of the mud, rocks, or submerged plant
debris. Smaller animals and algae that live on firmer
substrates are sometimes called aufwuchs. In addition
to the attached aufwuchs, macrophytes support some
larger, motile invertebrates which either feed on the
aufwuchs (e.g., grazing snails; Chapter 10) or use the
plant stems and leaves as perching sites for resting or
foraging (e.g., sprawling dragonfly nymphs; Chapters
17 and 18).
C. Wetlands, Ephemeral Ponds, and Swamps
For many years the public has ignored or labeled as
undesirable the vast acreage of wetlands, ephemeral
ponds, and swamps in North America, considering
them breeding grounds for mosquitoes and snakes.
Many have been drained, filled, and bulldozed for housing and commercial development without regard to
their intrinsic value to wildlife and the environment.
Our knowledge of these fascinating aquatic ecosystems
is relatively limited in comparison to information available about lakes and streams. Interestingly enough, one
factor that has begun reversing this trend has been the
implementation of strict environmental laws to protect
the broad category of wetlands. Such laws have also led
to an influx of research funds to study these ecosystems
which have spawned a huge variety of journal articles
(e.g., Zimmer et al., 2000), books (e.g., Batzer et al.,
1999; Mitsch and Gosselink, 2000), and even a new
journal (Wetlands).
In its broader definition, the term freshwater
wetlands refers to nontidal ecosystems whose soils are
saturated with water on a permanent or seasonal basis.
Emergent aquatic vegetation is prominent and may
alternate with annual terrestrial plants in these marshy,
or palustrine, habitats. The extensive biomass of trees,
herbaceous vegetation, grasses, and other plants in
many wetlands is responsible for their recently recognized abilities to help filter pollutants from the environment. Wetlands vary from shallow, seasonally
ephemeral ponds (such as Carolina bays and other
pocosins, Sharitz and Gibbons, 1982) to relatively
permanent vegetation-choked marshes to semi-lotic
alluvial swamps (Fig. 10; see also Mitsch and Gosselink, 2000).
Because wetlands are generally more ephemeral
than other lentic ecosystems, their biotic communities
are strongly influenced by water cycles. Wetlands invertebrates are affected by ecosystem permanency (e.g.,
years since last exposure), predictability of drying, and
season of drying (if at all). The nature of the community also reflects volume and depth of open water and
sometimes current velocity (for alluvial swamps). See
Batzer et al. (1999) for habitat specific information on
invertebrate ecology from most types of freshwater
wetlands in North America.
Invertebrates of seasonal wetlands, such as the Carolina bays of the southeastern United States, have
evolved various characteristics that have directly or
indirectly adapted them for ecosystems that alternate
between aquatic and terrestrial states. Wiggins et al.
(1980) divided animals of temporary pools into four
groups based on their adaptations for tolerating or
avoiding drought and their period of recruitment.
“Group 1 are year-round residents incapable of active
dispersal, which avoid desiccation either as resistant
stages or by burrowing into pool sediments. Group 2
are spring recruits which must oviposit on water but
subsequently aestivate and overwinter in the dry basin
in various stages. Group 3 are summer recruits ovipositing in the dry basin and overwintering as eggs or larvae.
Group 4 are nonwintering migrants leaving the pool before the dry phase, which is spent in permanent water;
they return in spring to breed.” When a temporary pool
refills after a drought, the first colonizers are usually
detritivores which exploit the abundant biomass of recently dead and decaying terrestrial vegetation; these
species provide the animal tissue that supports the later
arrival of predaceous invertebrates (Wiggins et al.,
1980). Many species in ephemeral environments are
ecological generalists, because they live in both temporary and permanent aquatic ecosystems.
Alluvial swamps are at an opposite continuum from
temporary wetland pools. Contrasting with the typical
picture of a swamp as a stagnant marsh, alluvial (or
riverine) swamps contain many areas of slowly to rapidly
moving waters; portions of these wetlands somewhat resemble a highly braided stream. While it is true that
southeastern alluvial swamps, such as those bordering
the Savannah River between South Carolina and Georgia, are infested with alligators and poisonous snakes,
they are otherwise extremely beautiful ecosystems with
very few aerial insect pests once you have moved inward
from the surrounding brush and terrestrial forest. Alluvial swamps are extremely heterogeneous, organically
rich environments; oxygen does not appear to be limiting, at least in the shaded, flowing water regions.
2. An Overview of Freshwater Habitats
A
B
FIGURE 10
(A) Photograph of a low-flow portion of a bald cypress — water tupelo, alluvial swamp of
the Savannah River. Floating platforms in center are suspending wood snags as part of a colonization
study by Thorp et al. (1995). (B) Small wetland area in northern New York created by a beaver dam.
Shown are cattails (foreground), open water covered by duckweed (middle), and various emergent vegetation and woody plants (photographs by J. H. Thorp).
37
38
J. H. Thorp and A. P. Covich
The many soft- and hard-sediment habitats of
swamps are home to a great diversity and density of
aquatic invertebrates. In a study of an alluvial swamp
in South Carolina, Thorp et al. (1985) showed that invertebrates rapidly colonized wood snags, reaching a
rough steady state in numbers within the first week of
an 8-week study. Peak densities were equivalent to
nearly 18,000 animals/m2! “Filter-feeding taxa were
numerically dominant early but soon were subordinate
to gatherer and scraper functional feeding groups.
Current velocity, seston particle size [as a food resource], dispersal capacity and competition for space
may be important factors affecting community structure and colonization patterns in these aquatic ecosystems” (p. 56).
D. Hypersaline Lakes
Natural saline lakes are common worldwide but
generally have neither the average size nor abundance
of freshwater lakes (Eugster and Hardie, 1978; Hammer, 1984). They are frequently encountered in the
prairies and the Great Basin of the western United
States and in the provinces of Alberta, Saskatchewan,
and British Columbia of western Canada (Hammer,
1984). The Great Salt Lake of Utah is the largest saline
lake in North America. Saline lakes usually occur in
more arid regions where closed basins promote high
concentrations of salts (Fig. 11). These lentic ecosystems include saltern lakes, which are high in sodium
chloride; lakes with large concentrations of sulfates and
borates; and soda lakes characterized by abundant
sodium carbonates and bicarbonates. Salinities in the
Great Salt Lake have been recorded as high as 200 g/L
(versus an average of 35 g/L in the world’s oceans). In
addition to high salinities, hypersaline lakes differ in
pH, metal concentrations, and alkalinity from more
typical inland lakes.
The abiotic characteristics of saline lakes make
them extremely rigorous environments. Cyanobacteria
colonize saline lakes as do some sedges, but the submerged macrophyte flora is relatively sparse in comparison to that in freshwater lakes. Because invertebrate
density and diversity are enhanced by abundant littoral
plants, the depauperate flora of saline lakes and the
lack of certain other microhabitats undoubtedly depress the invertebrate fauna of briny pools. Few invertebrates of inland waters can regulate osmotic and
ionic concentrations of their tissues in a hypertonic environment. As a result, hypersaline environments contain very few species, although their densities may be
high. For example, the only permanent metazoa in
Mono Lake, California (Fig. 11), are the ubiquitous
brine shrimp, Artemia salina, and the brine fly genus
Ephydra (see also Herbst, 1999). High densities of
brine shrimp in Mono Lake are crucial to survival of
FIGURE 11 Saline Mono Lake in arid central California. The light “rocks” in the foreground are
actually calcium carbonate mounds known as “tufa”; many of these arise within the lake proper. Visible
on the island in the background is a small volcanic cone (photograph by J.H. Thorp).
2. An Overview of Freshwater Habitats
migratory eared grebes, Podiceps nigricollis, which
molt flight feathers while at Mono Lake, depending on
these grazing crustaceans as their sole food source
(Cooper et al., 1984). This simple but important food
chain is threatened by the possible demise of the brine
shrimp as a result of rising salinities brought on by removal of freshwater from the Mono Lake catchment to
provide water for the human population in Southern
California. Other saline lakes support more species, including the rotifer Brachionus plicatilis, the calanoid
copepod Diaptomus nevadensis, and several families of
flies, beetles, and true bugs.
As saline lakes become less salty, the diversity and
density of their flora and fauna are enhanced. For example, the meromictic Waldsea Lake of Saskatchewan,
which is one of the better studied saline ( 3 ppt total
dissolved solids) inland lakes in Canada, has a much
higher species diversity than the much saltier Mono
Lake. It contains two macrophyte species, some filamentous algae, a sparse phytoplankton population,
and a dense but rather uniform population of zooplankton, dominated by the calanoid copepod Diaptomus connexus, the rotifers Hexarthra fennica and Brachionus plicatilis, and the water flea Daphnia similis
(Hammer, 1984). The littoral zone supports high numbers of relatively few species, including several species
of true bugs, nine genera of beetles, midges (mostly
Cricotopus), a few caddisflies, the damselfly Enallagma
clausum, one snail species (Lymnaea stagnalis), and
several genera of crustaceans (see references in Hammer, 1984). Although Waldsea Lake is certainly diverse
compared to the hypersaline Mono Lake, its community is depauperate in comparison to most permanent,
freshwater lentic environments.
The inland waters of North America are sufficiently diverse and abundant to prevent the valid formation of any simple, all-encompassing theory on how
biotic communities of these aquatic ecosystems are regulated. This does not imply, however, that we should
abandon the task of assimilating and evaluating data
from a diversity of ecosystems in order to understand
better the effects of a few identifiable factors, such as
the influence of habitat characteristics. A comparative
approach, combined with intensive descriptive and
experimental studies, will in the long run enhance our
ability to develop a holistic view of stream and lake
ecosystems. As we have shown in this chapter and as
you will learn in Chapters 3 – 23, habitat preferences of
an organism are extremely crucial in determining its
distribution, survival, and reproductive output. The nature of the habitat, however, is only one of a large suite
of factors influencing distribution and density of freshwater invertebrates.
39
LITERATURE CITED
Allan, J. D. 1995. Stream ecology: Structure and function of running
waters. Chapman and Hall, New York.
Barnby, M. A., Resh, V. H. 1988. Factors affecting the distribution
of an endemic and a widespread species of brinefly (Diptera:
Ephydridae) in a northern California thermal saline spring. Annals of the Entomological Society of America 81:437 – 446.
Batzer, D. P., Rader, R. B., Wissinger, S. A. 1999. Invertebrates in
freshwater wetlands of North America: ecology and management, Wiley, New York, 1100 p.
Benke, A. C., Jacobi, D. I. 1994. Production dynamics and resource
utilization of snag-dwelling mayflies in a blackwater stream.
Ecology 75(5):1219 – 1232.
Benke, A. C., Wallace, J. B. 1997. Trophic basis of production
among riverine caddisflies: implications for food web analysis.
Ecology 78:1132 – 1145.
Boulton, A. J., Findlay, S., Marmonier, P., Stanley, E. H., Valett,. H.
M. 1998. The functional significance of the hyporheic zone in
streams and rivers. Annual Review of Ecology and Systematics
29:59 – 81.
Brunke, M., Gonser, T. 1999. Hyporheic invertebrates — the clinal
nature of the interstitial communities structured by hydrological
exchange and environmental gradients. Journal of the North
American Benthological Society 18:344 – 362.
Calow, P., Petts, G. E. Eds. 1992. The rivers handbook: hydrological
and ecological principles, Vol. 1, Blackwell Science, Oxford,
England, 526 p.
Carpenter, S. R. Ed. 1988. Complex interactions in lake communities, Springer-Verlag, New York. 283 p.
Cole, G. A. 1994. Textbook of limnology, 4th ed. Mosby, St. Louis,
MO, 412 p.
Connell, J. H. 1978. Diversity in tropical rain forests and coral reefs.
Science 199:1302 – 1310.
Cooper, S. D., Winkler, D. W., Lenz, P. H. 1984. The effects of grebe
predation on a brine shrimp population. Journal of Animal
Ecology 53:51 – 64.
Covich, A. P. 1988. Geographical and historical comparisons of
neotropical streams: biotic diversity and detrital processing in
highly variable habitats. Journal of the North American Benthological Society 7:361 – 386.
Culver, D. C., Master, L. L., Christman, M. C., Hobbs, H. H. 2000.
Obligate cave fauna of the 48 contiguous Unites States. Conservation Biology 14:386 – 401.
Delucchi, C. M., Peckarsky, B. A. 1989. Life history patterns of insects in an intermittent and a permanent stream. Journal of the
North American Benthological Society 8:308 – 321.
Eugster, H. P., Hardie, L. A. 1978. Saline lakes, in: A. Lerman, Ed.,
Lakes: chemistry, geology, and physics. Springer-Verlag, New
York, pp. 237 – 293.
Forester, G. E., Dudley, T. L., Grimm, N. B. 1999. Trophic interactions in open systems: effects of predators and nutrients on
stream food chains. Limnology and Oceanography
44:1187 – 1197.
Garton, D. W., Haag, W. R. 1993. Seasonal reproductive cycles and
settlement patterns of Dreissena polymorpha in western Lake
Erie. Chapter 6, in: Nalepa, T.F. and D.W. Schloesser, Eds.,
Zebra mussels: biology, impacts, and control. Lewis Publishers,
Boca Raton, FL, pp. 111 – 128.
Gayraud, S., Philippe, M., Maridet, L. 2000. The response of benthic
macroinvertebrates to artificial disturbance: drift or vertical
movement in the gravel bed of two sub-alpine streams? Archiv
für Hydrobiologie 147:431 – 446.
40
J. H. Thorp and A. P. Covich
Gibert, J., Danielopol, D. L., Stanford, J. A. Eds. 1994. Groundwater
ecology. Academic Press, San Diego, 571 p.
Gordon, N. D., McMahon, T. A., Finlayson, B. L. 1992. Stream hydrology: an introduction for ecologists. Wiley, New York, 526 p.
Hammer, U. T. 1984. The saline lakes of canada, in: F.B. Taub, Ed.,
Ecosystems of the world, No. 23: lakes and reservoirs. Elsevier,
New York, pp. 521 – 540.
Herbst, D. B. 1999. Biogeography and physiological adaptations of
the brine fly genus Ephydra (Diptera: Ephydridae) in saline waters of the Great Basin. Great Basin Naturalist 59:127 – 135.
Hershey, A. E. 1985. Effects of predatory sculpin on the chironomid
communities in an arctic lake. Ecology 66:1131 – 1138.
Howarth, R. W., Schneider, R., Swaney, D. 1996. Metabolism and
organic carbon fluxes in the tidal freshwater Hudson River.
Estuaries 19:848 – 865.
Hutchinson, G. E. 1957. A treatise on limnology. I. Geography,
physics, and chemistry. Wiley, New York, 1015 p.
Hutchinson, G. E. 1967. A treatise on limnology. II. Introduction to
lake biology and limnoplankton. Wiley, New York, 1115 p.
Hutchinson, G. E. 1975. A treatise on limnology. III. Limnological
botany. John Wiley, New York, 660 p.
Hutchinson, G. E. 1993. A treatise on limnology. IV. The zoobenthos. Wiley, New York, 944 p.
Hynes, H. B. N. 1970. The ecology of running waters. University of
Toronto Press, Toronto, Ont. 555 p.
Hynes, H. B. N. 1989. Keynote address, in: Dodge, D. P., Ed., Proceedings of the International Large River Symposium, Can. Sp.
Publ. Fish. Aquat. Sci. 106, pp. 5 – 10.
Jack, J. D., Thorp, J. H. 2000. Field experiments on regulation of
potamoplankton in a large river by a benthic suspension-feeding
mussel, Dreissena polymorpha. Freshwater Biology (in press).
Jones, J. B., Mulholland, P. J. Eds. 2000. Streams and ground waters.
Academic Press, San Diego, 425 p.
Junk, W. J., Bayley, P. B., Sparks, R. E. 1989. The flood pulse concept
in river-floodplain systems, in: Dodge, D. P., Ed., Proceedings of
the International Large River Symposium, Can. Sp. Publ. Fish.
Aquat. Sci. 106, pp. 110 – 127.
Lampert, W., Sommer, U. 1997. Oxford University Press, New York,
382 p. [Translation of a German text published in 1993.]
Ledger, D. C. 1981. The velocity of the River Tweed and its tributaries. Freshwater Biology 11:1 – 10.
Leopold, L. B. 1953. Downstream change of velocity in rivers. American Journal of Science 251: 606 – 624.
Longley, G. 1986. The biota of the Edwards aquifer and the implications for paleozoogeography. in: Abbott P. L., Woodruff C.M.,
Jr., Eds., The Balcones Escarpmernt, Geological Society of
America, Central Texas, pp. 51 – 54.
Malard, F., Hervant, F. 1999. Oxygen supply and the adaptations of
animals in groundwater. Freshwater Biology 41:1 – 30.
Matthews, W. J. 1988. North American prairie streams as systems
for ecological study. Journal of the North American Benthological Society 7:387 – 409.
Merritt, R. W., Cummins, K. W. Eds. 1996. An introduction to the
aquatic insects of North America. 3rd ed., Kendall/Hunt,
Dubuque, Iowa.
Meyer, J. L. 1990. A blackwater perspective on riverine ecosystems.
BioScience 40:643 – 651.
Mihuc, T. B. 1997. The functional trophic role of lotic primary consumers: generalist versus specialist strategies. Freshwater Biology 37:455 – 462.
Minckley, W. L. 1963. The ecology of a spring stream Doe Run,
Meade County, Kentucky. Wildlife Monographs No. 11 (A publication of the Wildlife Society), 124 p.
Minshall, G. W. 1988. Stream ecosystem theory: a global perspective.
Journal of the North American Benthological Society. 7:263 – 288.
Mitsch, W. J., Gosselink, J. G. Eds. 2000. Wetlands, 3rd ed., Wiley,
New York, 750 p.
Naiman, R. J., Melillo, J. M., Hobbie, J. E. 1986. Ecosystem alteration of boreal forest streams by beaver (Castor canadensis).
Ecology 67:1254 – 1269.
Newbury, R. W. 1984. Hydrologic determinants of aquatic insect
habitats, in: Resh, V. H., Rosenberg D. M., Eds., The ecology of
aquatic insects. Praeger, New York, pp. 323 – 357.
Pace, M. L., Findlay, S. E. G., Fischer, D. 1998. Effects of an invasive
bivalve on the zooplankton community of the Hudson River.
Freshwater Biology 39:103 – 116.
Pace, M. L., Findlay, S. E. G., Lints, D. 1992. Zooplankton in advective environments: The Hudson River community and a comparative analysis. Canadian Journal of Fisheries and Aquatic
Sciences 49:1060 – 1069.
Reckendorfer, W., Keckeis, H., Winkler, G., Schiemer F. 1999. Zooplankton abundance in the River Danube, Austria: the significance of inshore retention. Freshwater Biology 41:583 – 591.
Resh, V. H., Brown, A. V., Covich, A. P., Gurtz, M. E., Li, H. W., Minshall, G. W., Reice, S. R., Sheldon, A. L., Wallace, J. B., Wissmar,
R. C. 1988. The role of disturbance in stream ecology. Journal of
the North American Benthological Society 7:433 – 455.
Ritter, D. F. 1986. Process geomorphology, 2nd ed., Wm. C. Brown,
Debugue, Iowa.
Sedell, J. R., Richey, J. E., Swanson, F. J. 1989. The river continuum
concept: a basis for the expected behavior of very large rivers?
in: Dodge, D.P., Ed., Proceedings of the International Large
River Symposium, Can. Sp. Publ. Fish. Aquat. Sci. 106, pp.
49 – 55.
Sharitz, R. R., Gibbons, J. W. 1982. The ecology of southeastern
shrub bogs (pocosins) and Carolina bays: a community profile.
FWS/OBS-82/04. U.S. Fish and Wildlife Service, Division of
Biological Services, Washington, D. C, 93 p.
Stanford, J. A., Hauer, F. R., Ward, J. V. 1988. Serial discontinuity in
a large river system. Verhandlungen Internationale Vereinigung
fur theoretische und angewandte Limnologie 23:114 – 118.
Stanford, J. A., Ward, J. V. 1988. The hyporheic habitat of river
ecosystems. Nature 335:64 – 66.
Statzner, B., Gore, J. A. Resh, V. H. 1988. Hydraulic stream ecology:
observed patterns and potential applications. Journal of the
North American Benthological Society 7:307 – 360.
Statzner, B., Higler, B. 1985. Questions and comments on the River
Continuum Concept. Canadian Journal of Fisheries and
Aquatic Sciences 42:1038 – 1044.
Strayer, D. 1985. The benthic micrometazoans of Mirror Lake, New
Hampshire. Archive für Hydrobiologie/Suppl. 72(3):287 – 426.
Strayer, D. 1986. The size structure of a lacustrine zoo-benthic community. Oecologia 69:513 – 516.
Strayer, D., May, S. E., Nielsen, P., Wollheim, W., Hausam, S. 1995.
An endemic groundwater fauna in unglaciated eastern North
America. Canadian Journal of Zoology 73:502 – 508.
Strayer, D. L., Reid, J. W. 1999. Distribution of hyporheic cyclopoids
(Crustacea: Copepoda) in the eastern United States. Archiv für
Hydrobiologie 145:79 – 92.
Strahler, A. N. 1952. Dynamic basis of geomorphology. Geological
Society of America Bulletin 63:1117 – 1142.
Strenth, N. E., Norton, J. D., Longley, G. 1988. The larval development of the subterranean shrimp Palaemonetes antrorum Benedict (Decapoda, Palaemonidae) from central Texas. Stygologia
4:363 – 370.
Sweeney, B. W., Vannote, R. L. 1978. Size variation and the distribution of hemimetabolous aquatic insects: two thermal equilibrial
hypotheses. Science 200:444 – 446.
Taub, F. B. Ed. 1984. Lakes and reservoirs. Ecosystems of the world,
Vol. 23, Elsevier, New York, 643 p.
2. An Overview of Freshwater Habitats
Thorp, J. H., Cothran, M. L. 1984. Regulation of freshwater community structure at multiple intensities of dragonfly predation.
Ecology 65:1546 – 1555.
Thorp, J. H., Delong, M. D. 1994. The riverine productivity model:
an heuristic view of carbon sources and organic processing in
large river ecosystems. Oikos 70(2):305 – 308.
Thorp, J. H., Delong, M. D., Casper, A. F. 1998. In situ experiments
on predatory regulation of a bivalve mollusc (Dreissena polymorpha) in the Mississippi and Ohio Rivers. Freshwater Biology 39:649 – 661.
Thorp, J. H., Delong, M. D., Greenwood, K. S., Casper, A. F. 1998.
Isotopic analysis of three food web theories in constricted and
floodplain regions of a large river. Oecologia 117:551 – 563.
Thorp, J. H., McEwan, E. M., Flynn, M. F., Hauer, F. R. 1985. Invertebrate colonization of submerged wood in a cypress-tupelo swamp
and blackwater stream. American Midland Naturalist 113:56–68.
Vannote, R. L., Minshall, G. W., Cummins, K. W., Sedell, J. R.,
Cushing, C. E. 1980. The river continuum concept. Canadian
Journal of Fisheries and Aquatic Sciences 37:130 – 137.
Ward, J. V. 1986. Altitudinal zonation in a Rocky Mountain stream.
Archiv fur Hydrobiologie, Suppl. 74:133 – 199.
Ward, J. V. 1989. The four-dimensional nature of lotic ecosystems.
Journal of the North American Benthological Society 8:2 – 8.
Ward, J. E., Standford, J. A. 1983. The intermediate disturbance hypothesis: an explanation for biotic diversity patterns in lotic
41
ecosystems, in: Fontaine T.D., Bartell, S.M. Eds., Dynamics of
lotic ecosystems. Ann Arbor Science Publishers, Ann Arbor, MI,
pp. 347 – 356.
Wetzel, R. G. 1983. Limnology, 2nd ed., Saunders, Philadelphia,
857 p.
Wetzel, R. G. 2001. Limnology, 3rd ed., Academic Press, San Diego
(in press).
Wiggins, G. B., Mackay, R. J., Smith, I. M. 1980. Evolutionary and
ecological strategies of animals in annual temporary pools.
Archive fur Hydrobiologie/Suppl. 58(1):97 – 206.
Williams, D. D. 1984. The hyporheic zone as a habitat for aquatic
insects and associated arthropods. in: Resh, V.H., Rosenberg,
D.M. Eds., The ecology of aquatic insects. Praeger, New York,
pp. 430 – 455.
Williamson, C. E., Hargreaves, B., Orr, P. S., Lovera, P. A. 1999.
Does UV play a role in changes in predation and zooplankton
community structure in acidified lakes? Limnol. Oceanogr.
44(3, part 2):774 – 783.
Winterbourn, M. J. 1982. The River Continuum Concept-reply to
Barmuta and Lake. New Zealand Journal of Marine and Freshwater Research 16:229 – 231.
Zimmer, K. D., Hanson, M. A., Butler, M. G. 2000. Factors influencing invertebrate communities in prairie wetlands: a multivariate
approach. Canadian Journal of Fisheries and Aquatic Sciences
57:76 – 85.
This Page Intentionally Left Blank
3
PROTOZOA
William D. Taylor
Robert W. Sanders
Department of Biology
University of Waterloo
Waterloo, Ontario N2L 3G1, Canada
Department of Biology
Temple University
Philadelphia, Pennsylvania 191ZZ
I. Introduction
II. Anatomy and Physiology
A. Protozoa as Cells and Organisms
B. Protozoan Organelles
C. Anatomy of Flagellates
D. Anatomy of Amoeboid Protozoa
E. Anatomy of Heliozoa
F. Anatomy of Ciliates
G. Environmental Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Collecting, Rearing, and Preparation
for Identification
I. INTRODUCTION
Protozoa are ubiquitous; they are present in an
active state in all aquatic or moist environments, and
cysts are present everywhere in the biosphere, ready to
give rise to active populations. They play an important
role in many natural communities, although they are
often overlooked. In the laboratory, protozoa often play
the role of model organisms; cell biologists, physiologists, geneticists, and even developmental biologists frequently turn to protozoa to address questions that
would be more difficult to answer with metazoa, but in
the hope that the answers are relevant to metazoa. The
result is a surprising amount of information about a relatively few species. Accordingly, detailed monographs are
available on several taxa, including Blepharisma (Giese,
1973), Paramecium (Wichterman, 1986), Tetrahymena
(Elliott, 1973) and Amoeba (Jeon, 1973). Each monograph contains thousands of references. Ecologists also
have used protozoan populations and communities as
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
V. Identification of Protozoa
A. Notes about the Key
B. Taxonomic Key to Functional Groups
of Protozoa
C. Taxonomic Key to Major Groups
and Common Genera of Flagellated
Protozoa
D. Taxonomic Key to the Major Groups
and Common Genera of Protozoa
with Pseudopodia
E. Taxonomic Key to Major Groups
and Common Genera of Ciliates
Literature Cited
model systems to investigate processes, such as competition and predation, or to test models, such as island
biogeography, that would be much more difficult to
study using larger organisms. Again, the result has been
an extensive body of literature that is largely restricted to
a few species and/or to unusual or artificial habitats.
In the last 20 years, there has been an explosion of
information about the ecology of free-living protozoa,
especially the planktonic species that were virtually ignored in the earlier literature. This increase in interest
followed the development of direct-count methodologies for aquatic bacteria, and an appreciation of the importance of bacteria and their predators in aquatic systems. The first focus was on the heterotrophic
flagellates and ciliates that consumed bacterial production and passed it to larger consumers, the “microbial
loop.” As aquatic ecologists came to realize that protozoa are not only present, but play a major quantitative
role in material and energy flow, this interest expanded
to other feeding guilds and freshwater environments.
43
44
W. D. Taylor and R. W. Sanders
The goal of this chapter is to provide an introduction to the ecology and classification of protozoa for
ecologists who wish to include them in studies of
aquatic environments. The chapter focuses on the biology of the animal-like protozoa, largely ignoring the
related pigmented organisms which are covered in
phycological sources. The chapter is oriented toward
the organisms, not the freshwater habitats themselves.
Readers wishing to learn about the fauna of particular
habitats might consult the bibliography on freshwater
protozoa prepared by Finlay and Ochsenbein-Gattlen
(1982) or some more recent works on particular habitats: lake plankton (Beaver and Crisman, 1989;
Laybourn-Parry, 1992; Riemann and Christoffersen,
1993; Carrias et al., 1996 Foissner et al. 1999),
streams (Bott and Kaplan, 1990; Carlough and Meyers,
1991; Sleigh et al., 1993), and wetlands (Pratt and
Cairns, 1985; Decamp and Warren, 1999).
What are protozoa?
Protozoa are unicellular or colonial eukaryotes, including all of the heterotrophic and motile ones. It is a
taxon of convenience. Although the protozoa were
considered to be a single phylum of eukaryotic animals
in early classifications, modern treatments distribute
the protozoa among many phyla. This reflects the immense diversity of this assemblage, and the fact that
differences among these unicellular organisms are at
least as profound as those which separate plant and
animal phyla (see Section III.D). Margulis and
Schwartz (1988), in their five-kingdom classification of
living organisms, place the unicellular eukaryotes and
some of their multicellular descendants, 27 phyla in
all, in the kingdom Protoctista. Other modern classifications partition the protists into more or fewer phyla.
The classification of unicellular eukaryotes is in a state
of flux, with much work to be done. In any event, it is
among these groups that the organisms we know as
protozoa are scattered.
For the purpose of this chapter, we define the term
protozoa as those unicellular or colonial eukaryotes
that are heterotrophic, and will restrict ourselves to a
discussion of free-living, phagotrophic forms in freshwater. Among the protozoan phyla we will not discuss
here are the marine Foraminifera and Radiolaria, the
soil-dwelling Labyrinthulamycota, Plasmodiophoromycota and Acrasida, the saprophytic Hyphochytridiomycota, Oomycota and Chytridiomycota, and the parasitic Apicomplexa, Microspora, and Myxospora. The
classification we use, and citations to figures illustrating
the major taxa, are outlined in Table I. It is designed to
be as current as possible, but still retain the older
names. Therefore, it emphasizes classes, subclasses, or
orders, largely according to which retains the most
familiar name for the group in question. Our major
TABLE I Major Taxomomic Categories of Protozoa as Used
in This Chapter
Functional Group
Phylum
Class
subclass
Order
Phytoflagellates (largely autotrophic,
but some mixotrophic and heterotrophic)
Cryptophyta
Dinoflagellata
Euglenida
Chrysophyta
Chlorophyta
Zooflagellates (heterotrophic)
Zoomastigina
Choanoflagellida
Kinetoplastida
bicoecids
diplomonads
cercomonads
Ameboid Protozoa
Karyoblastea
Rhizopoda
Heterolobosea
Schizopyrenida
Lobosea
Gymnamoebia
Euamoebida
Leptomyxida
Acanthopodida
Testacealobosia
Arcellinida
Himatismenida
Filosea
Aconchulinida
Testaceafilosida
Granuloreticulosida
Athalamea
Monothalamea
Foraminiferea (marine only)
Actinopoda
Heliozoa
Actinophryida
Desmothoracida
Ciliophryida
Centrohelida
Rotosphaerida
Ciliated Protozoa
Ciliophora
Karyorelictea
Heterotrichea
Spirotrichea
Hypotrichia
Choreotrichia
Stichotrichia
Oligotrichia
Odontostomatida
Armophorida
Litostomatea
Figure reference
13J
13A – C
13D, F – I
13K – N; 14A
13E
14B – H
14J, L – P
14I
14M
14K
16V
15A – C
15D – M,O,P
15N
16B – Q
16A
16R, 17A
16S – W; 17B – K
17P – R
17L – O
18B,C
18A
18E
18D,F,H
18G
23J
22A,B,D – F
20X,Y
21S,T,U,X
21A – I,N – Q
21V,W
21,J,M,L
21K,R
22G,I,K,L,N,P – W;
23D – I
(Continues)
3. Protozoa
45
TABLE I (Continued)
Functional Group
Phylum
Class
subclass
Order
Phyllopharyngea
Phyllopharyngia
Suctoria
Nassophore
Nassulida
Microthoracida
Colpodea
Prostomatea
Prostomatida
Prorodontidae
Plagiopylea
Oligohymenophorea
Peniculia
Scuticociliatia
Hymenostomatia
Peritrichia
Figure reference
24T
19A – P
20Z
20V,W
22C; 23K; 24N,P – S
22H,J,O
23A – C
24M
24A,B,D,E,G,I,J
23L – X
24 C,F,H,O
20 A – U
points of reference were Page (1998), Lynn and Small
(1997), and various chapters in Lee et al. (1985) and
Margulis et al. (1989).
Despite the fact that the term protozoa is perhaps
not a proper taxonomic group, it remains a useful
functional one. For ecologists conducting studies involving aquatic animals, including those of microscopic
proportions, it is natural to include the animal-like
protozoa in those studies. After all, it is functional similarity that generally concerns the practising zoologist
or ecologist.
II. ANATOMY AND PHYSIOLOGY
A. Protozoa as Cells and Organisms
Large protozoa that have many nuclei, or large,
amitotic, polyploid nuclei, may be best thought of as
acellular rather than unicellular creatures. Nonetheless,
some species have been studied extensively by cell
biologists as model cells, and ecologists interested in
protozoa can glean much useful information from their
labors. This section will rely heavily on information
from those few taxa. We will first describe some of the
organelles which are important and widely distributed
among protozoa, then discuss major groups of protozoa with respect to their more unique organelles.
Lastly, we will revert to an overall view in discussing
environmental physiology of protozoa. In all sections,
we will emphasize those organelles that relate to
FIGURE 1
The formation and fate of food vacuoles, autophagic
vacuoles, and pinocytotic vacuoles in Tetrahymena pyriformis. (From
Elliott and Clemmons, 1966.)
feeding, locomotion, ecology, and morphology at the
light-microscope level.
B. Protozoan Organelles
1. Food Vacuoles
Phagocytosis in protozoa leads to the formation of
food vacuoles (Fig. 1). Digestion is, as one would expect, primarily intracellular in free-living protozoa. The
formation and behavior of food vacuoles is best known
in the ciliates Tetrahymena (see Rasmussen, 1976;
Nilsson, 1976, 1987) and Paramecium (Allen, 1984). A
more general review is provided by Nisbet (1984).
The formation of food vacuoles is stimulated by
the presence of particulate food. In protein broth
medium, Tetrahymena shows very slow rates of food
vacuole formation, ingestion, and growth unless
particles are present (Ricketts, 1972), even though it
can grow without forming food vacuoles in complex
media via carrier-mediated uptake of dissolved
compounds across the plasma membrane. During
starvation, vacuoles resembling food vacuoles may
be created to digest cellular components. Formation
of such “cytolosomes” is also known from Amoeba
(Chapman-Andresen, 1973).
46
W. D. Taylor and R. W. Sanders
Many protozoa have a fixed site of food-vacuole
formation; a cytostome with accompanying structures
to aid in ingestion. In others, for example, Rhizopoda
and Heliozoa, the site of ingestion is flexible, perhaps
only limited to a characteristic region of the cell. The
Suctoria have many sites for ingestion. The contents of
a single vacuole may vary from a single large food item
in macrophagous species to thousands of small items,
such as bacteria, in microphagous forms. Similarly, individual protozoa may have one-to-many food vacuoles. The vacuoles of microphagous forms may be
perfectly spherical, while larger vacuoles containing
single prey may be quite irregular, conforming to the
contours of the prey. Macrophagous protozoa may distend themselves to surround large food items, leave
them protruding from their cytoplasm, or even share a
prey item among several individuals. Large food items
may be digested by cytoplasmic extensions in dinoflagellates and possibly other protozoa (Fig. 2).
Digestion may be initiated by the toxicysts of some
predatory ciliates. In others, it begins when granules
(acidosomes) fuse with the developing food vacuole.
The pH of the new vacuole drops precipitously, then
increases minutes later. Hydrolytic enzymes, such as
FIGURE 2 Phagocytosis of large food items by protoplasmic
extensions in the dinoflagellate Ceratium hirudinella (A – C), and the
consumption of a large diatom by a Paraphysomonas-like chrysophyte (D). (A – C from Hofeneder, 1930: D after Suttle et al. 1986.)
acid phosphatase, are introduced as the food vacuole
fuses with lysosomes. The size of the food vacuole may
diminish, reflecting the absorption of water and nutrients, and vacuoles may coalesce. Some protozoa, such
as ciliates, have a fixed site of egestion or cytoproct.
When the contents of the food vacuole are released, the
membrane surrounding the vacuole is retained and
recycled (Allen, 1984). It is unlikely that a cellular
equivalent of a digestive track exists in protozoa. Nevertheless, it has been possible to estimate the feeding
rate of protozoa in the field from the rate at which
food vacuoles are eliminated (Goulder, 1972).
2. Contractile Vacuoles
These organelles are widely distributed in protists
and are almost universal in freshwater protozoa and
sponges. They are absent in dinoflagellates and are generally lacking in those protists with rigid cell walls
and/or from marine or endosymbiotic habitats. A large
body of evidence suggests that contractile vacuoles
eliminate water and, thereby, counteract the tendency
of naked protists to swell in freshwater (Patterson,
1980). The activity of the contractile vacuole is correlated with changes in the osmotic balance between the
cell and its medium, and a loss of contractile function
results in swelling of the cell. Of course, the occurrence
of contractile vacuoles in naked, freshwater protists
and their absence in most rigid-walled, marine or endosymbiotic ones also support this theory.
Associated with the contractile vacuole is a specialized cytoplasm, the spongiome, which collects the
solute to be expelled by contractile vacuole. The spongiome may contain vesicles or canals visible with the
light microscope, and therefore be of diagnostic value
(Fig. 3). Patterson (1980) recognized four types of contractile vacuole complexes in protozoa, with six different patterns of behavior. Permanent contractile vacuole
pores are present in ciliates and are readily visible in
silver-stained specimens. The contractile vacuole may
have a fixed location in some other groups, for example, in the gullet or reservoir of cryptomonad or euglenoid flagellates, but the fixed pores, if they exist, are
not visible.
Several important functional aspects of the contractile vacuole complex remain unknown; for example, the composition of the fluid expelled, and, therefore, the role of the contractile vacuole complex in salt
balance and excretion. It is also debatable whether
expulsion of contractile vacuole contents is active or
passive, and how the activity of the contractile vacuole
is controlled.
Dinoflagellates possess a functionally homologous
organelle, the pusule. This organelle consists of a single
3. Protozoa
47
FIGURE 3
Top views of three morphologies of contractile vacuoles found in protozoa, showing
their cycles of activity from left to right as seen with a light microscope. (A) A vacuole which forms
by the coalescing of smaller vesicles, as is common in Rhizopoda. Vacuole (B) has ampullae in fixed
locations, while vacuole (C) also has visible canals. (After Patterson, 1980.)
chamber or tubule associated with a flagellar opening,
and it is lined with vesicles (Dodge and Gruet, 1987).
3. Cilia and Flagella
Flagella, cilia, and derived structures are widely
distributed in eukaryotes, including protozoa. Their ultrastructural similarity, including the basic 9 pairs 2
pattern of microtubules in the shaft, or 9 triplets in the
base, indicates homology despite the great diversity in
form and function. Many animal sense organs contain
cilia, which emphasizes that they are probably intrinsically sensory as well as motile. Cilia or flagella which
are thought to be sensory occur in many protozoa, especially the “brosse” kinetids of prostomateans, and
the flagellum of phototactic euglenids.
It is well established that the motile force in cilia
and flagella is the sliding of microtubules in the shaft
against each other. Therefore, the active movement is
the bending of shaft and not the movement of shaft relative to the base. The bending of shaft takes on the appearance of a beat or stroke in short cilia or a wave in
long flagella.
Flagella usually occur in pairs and are relatively
long (often longer than the cell that carries them, and
reaching lengths of up to 50 m). A major functional
and morphological dichotomy among flagella is between those with and without mastigonemes (Fig. 4).
Mastigonemes are thin, hairlike projections perpendicular to the shaft of the flagellum. They are not visible
with light microscopy, but are functionally important.
A distally directed wave of bending in a smooth flagellum will push the cell in the opposite direction to
which the flagellum is pointed, as in animal sperm. The
same wave form in a flagellum with mastigonemes will
pull the cell in the direction in which the flagellum is
pointed. To understand the motility of protozoa, especially those using cilia and flagella, it is necessary to appreciate that the apparent viscosity of water increases
as the size of organisms moving through it decreases
(Holwill, 1974; Fenchel, 1987). To flagellates, water is
a liquid so viscous that they sink only very slowly and
they not glide from inertia.
The difference between cilia and flagella is one of
form and function. Cilia are generally short and
densely packed. They are organized in parallel rows
(kineties) to produce fields of cilia whose activities are
coordinated (Fig. 5A). The movement of adjacent somatic cilia is not quite synchronous; rather, each is
slightly out of phase with its neighbors, or metachronous. A field of cilia, therefore, moves in waves that
may or may not correspond in direction to the actual
power stroke of the cilia. Compound ciliary organelles
or polykinetids are restricted fields of cilia acting as a
single unit in locomotion, as in the cirri of hypotrichs
(Fig. 20Y), or feeding, as in the oral apparatus of hymenostomes (Fig. 5B). They may be relatively long
compared to other cilia and fused so that their compound nature is not obvious under light microscopy,
except as indicated by their diameter.
Ciliary movement is faster than flagellar movement, even accounting for ciliates generally being larger
than flagellates (Sleigh and Blake, 1977). Indeed,
within both these groups there is relatively little increase in absolute swimming speed with body size;
48
W. D. Taylor and R. W. Sanders
FIGURE 5 The distribution of cilia on Tetrahymena (A). The
compound ciliary organelles associated with the mouth (B) are used
to collect bacteria into the forming food vacuole. Somatic cilia beat
metachronously to propel the cell (C). (D) The path of a single cilium
viewed from the side, showing the power stroke and the recovery
stroke in different planes.
FIGURE 4
Different uses of flagella in two sessile bacterivores, and
two swimming forms. Arrows indicate movement of water relative to
the cells. Note that distally directed waves of beating produce opposite forces in smooth flagella and flagella with mastigonemes. (A) The
choanoflagellate Monosiga, modified from Fenchel (1982a); (B)
Actinomonas, which is probably a heliozoan, also modified from
Fenchel; (C) a dinoflagellate; (D) a chrysophyte.
flagellates average about 0.2 mm/s, while ciliates average about 1 mm/s.
4. Extrusomes
Extrusive organelles or extrusomes are membranebound organelles associated with the pellicle of protists
and containing material which can be ejected or
extruded from the cell. Most are visible at the lightmicroscope level, at least in the extruded state. They
are widespread in occurrence, and diverse in structure
and function; they are probably not homologous. The
function of many is in doubt (Haackbell et al., 1990).
However, following Hausman (1978) we will group
them according to their functional and structural similarity for discussion, omitting those restricted to marine
groups.
Spindle trichocysts of nassophorean ciliates, especially those of Paramecium, are well known. They are
abundantly distributed over the cortex (e.g., 8000 per
cell in Paramecium) and each may discharge a pointed
projectile on a proteinaceous shaft in response to
mechanical or physical stimulation. These are probably
effective deterrents to some potential predators
(Harumoto, 1994) although specialists, such as the
predatory ciliate Didinium, are undaunted (Fig. 6). Expulsion of the trichocyst is extremely rapid and is driven
by a change in the paracrystalline structure of protein in
the shaft of the trichocyst. The increase in length of the
shaft following expulsion is approximately eightfold.
Extrusomes are found in several flagellate groups;
most dinoflagellates have spindle trichocysts similar to
those of ciliates. Until recently, these groups were not
considered to be even remotely related. However, recent
studies of ribosomal RNA suggest they are less distant
than one might suppose based on gross morphology
(Gunderson et al., 1987). Ejectisomes are found in most
3. Protozoa
FIGURE 6 (A)
Capture of Paramecium by Didinium. Note the use of toxicysts by Didinium, and the
spindle trichocysts released by Paramecium in response. (B) The oral apparatus and zone of contact, as
revealed by electron microscopy. Note that there are two varieties of extrusomes involved in Didinium’s attack, and that the deciliation of Paramecium is part of the result. (C) The ultrastructure of
spindle trichocysts of Paramecium in the resting (left) and discharged (right) state. (D)Ultrastructure of
Didinium pexicysts (left) and toxicysts (right). Both are shown in resting and discharged states. c; Capsule. ci; cilia. olm; outer limiting membrane p; discharged pexicysts. pr; protuberance of Paramecium
cytoplasm. t; discharged toxicyst. tc; terminal cap. tr; discharged trichocyst. (B and D are from Wessenberg and Antipa, 1970; C is from Hausman, 1978.)
49
50
W. D. Taylor and R. W. Sanders
FIGURE 7
Examples of extrusomes in other protozoa. (A) Kinetocysts from the axopodia of the heliozoan Hetrophrys (from Lee et al., 1985). (B) Discobolocysts from the cell surface of the chrysophyte
Ochromonas, and the steps from formation (1) to discharge (5). (C) Ejectisomes from the gullet of
Cryptomonas. a; undischarged. b-d; discharging, showing uncoiling in two directions and rolling of the
uncoiled ejectisome into a tube. e; discharged ejectisome. (D) Haptocysts from the tentacles of a suctorian. Arrows indicate the location of the extrusomes to the right. Extrusomes in A – C are from
Hausman (1978). Haptocysts in D are drawn from Rudzinska (1965).
3. Protozoa
cryptomonads and in a few other phytoflagellates
(Fig 7). They are typically dimorphic, although of similar morphology; larger ejectisomes are associated with
the gullet, while smaller ones are external to the gullet
and near the anterior of the cell. Because some cryptophytes are heterotrophic, it is tempting to speculate that
the ejectisomes are used in food capture. However, the
gullet is not the site of phagotrophy, so an offensive role
of these extrusomes is doubtful. The mechanism of expulsion of ejectisomes is very different from that of
spindle trichocysts; the nonexpelled ejectisome is a tight
coil of ribbonlike material which uncoils and rolls into
a tube on expulsion. A similarly coiled secondary structure forms a pointed tip.
Discobolocysts are also extrusomes of flagellates,
in this case primarily of chrysophytes. They are almost
spherical, rather than spindle-shaped, but their expulsion is reminiscent of spindle trichocysts; the shaft of
the discobolocyst elongates, apparently through a conformational change in the material in the shaft (Fig. 7).
Although their function is unknown, it is possible that
they discourage phagotrophs.
Extrusomes are used for capturing food by predatory ciliates in the classes Prostomatea and Litostomatea, which have toxicysts, and Phyllopharyngea,
which have haptocysts. Toxicysts are extruded by eversion, and in doing so release material which appears to
be toxic to the prey. Several types of toxicysts are
known, and up to three kinds may be found within a
species. The use of toxicysts by Didinium in capturing
Paramecium is illustrated in Figure 6. Although the
toxicysts of Didinium are concentrated on the proboscis, they may be elsewhere in other ciliates. Loxophyllum (Fig. 23I) has them in clusters along its lateral
margin, while Actinobolina (Fig. 22G) has toxicysts on
tentacles that are widely distributed over the body.
Haptocysts are found in the tips of the feeding
tentacles of Suctoria (Fig. 19). Prey (mostly other ciliates)
which touch the feeding tentacles are caught and held by
the extrusion of the haptocysts (Fig. 7) which penetrate
the prey, inject substances (probably including hydrolytic
enzymes) and aid in the fusion of membranes of the
predator and prey. The cytoplasm of the prey is then
drawn into the suctorian through the feeding tentacles.
Some Heliozoa possess extrusomes of various
types, which may give the elongate pseudopodia (axoopodia) a bumpy appearance (Fig. 7). Kinetocysts resemble haptocysts or small trichocysts. Extrusomes in
Heliozoa also include mucocysts (see below) and
electron-dense bodies that also expel amorphous material onto the cell surface. All probably facilitate prey
adhesion and capture.
Mucocysts are widespread in ciliates and flagellates. They secrete an amorphous material onto the cell
51
surface that, at least in many species, is involved in cyst
formation. Tetrahymena can use mucocysts to create a
temporary capsule in response to introduction of a dye;
this is probably a defensive reaction, although it is not
known which other stimuli cause this response. Wolfe
(1988) suggested that mucocysts may be the protozoan’s first line of defense. Mucocysts may also be
involved in nutrition; the mucous secreted may be reingested after attracting dissolved compounds or flocculating small food items (see Section III.C). Euglenoid
flagellates have larger muciferous bodies in addition to
mucocysts, although some authors do not differentiate
between the two. These provide a continuous coat of
mucilage on the surface of the cell and, in some species,
participate in cyst formation, adhesion to the substratum, or stalk formation.
C. Anatomy of Flagellates
The term flagellates is applied to protists with oneto-many flagella. In free-living taxa, as opposed to parasitic species, the number of flagella is limited; Paramastix has two rows of 8 – 12 flagella, but most others
have 1 – 4 (usually 2). There are several groups of heterotrophic flagellates in freshwater: choanoflagellates,
kinetoplastids, diplomonads, cercomonads, and bicoecids. These are raised to phyla by some authors,
while bicoecids are occasionally classified with chrysophytes. Some Rhizopoda, the schizopyrenids or amoeboflagellates, also have flagella.
Other groups of flagellates contain mostly or
entirely autotrophic forms with chloroplasts. However,
many of the pigmented, autotrophic taxa are capable
of phagotrophy, a condition referred to as mixotrophy
(Sanders, 1991). Among these autotrophic and
mixotrophic groups there are nonpigmented, wholly heterotrophic species. The groups with many mixotrophic
or heterotrophic taxa include cryptophytes, chrysophytes, dinoflagellates, and euglenoid flagellates. These
groups are usually considered as phyla.
The functional significance of mixotrophy varies
widely. Different species use the ability to phagocytose
other cells to supplement photosynthesis in low light,
to gain access to nutrients, or to survive long periods in
the dark (Jones, 1997). In keeping with our desire to
cover the animal-like protists, we will survey heterotrophic and mixotrophic flagellates, ignoring primarily autotrophic groups. Pigmentation and chloroplast morphology are important taxonomic characters
for some groups, but here we will emphasize features
found in heterotrophs.
Choanoflagellates, or collared flagellates, are distinctive for the collar that surrounds the single flagellum (Figs. 4, 14C–H). They bear a strong resemblance
52
W. D. Taylor and R. W. Sanders
to choanocytes of Porifera. The collar may be difficult
to distinguish with light microscopy; but when examined with electron microscopy, it is evidently composed
of microvilli. The microvilli intercept bacteria drawn
past the cell by the flagellum. Because most choanoflagellates are attached or colonial, the distally directed
waves of the smooth flagellum serve to push water past
the protozoan. Bacteria caught on microvilli are transported to the cell and ingested by pseudopodia at the
base of the collar (Leadbeater and Morton, 1974).
Many choanoflagellates attach to the substrate, and
many have an external, loose-fitting covering or lorica
(although, again, it may be difficult to see with the light
microscope). In one marine family (Acanthoecidae), the
lorica is basketlike.
Bicoecids (Fig. 14I) resemble choanoflagellates, although they lack a collar. Like choanoflagellates, they
are enclosed in a lorica and have a flagellum that is
used to create a feeding current. A second flagellum lies
along the cell and continues posteriorly to become an
attachment to the base of the lorica.
Kinetoplastids (Fig. 14J,N – P) are known mostly as
parasites, especially Trypanosoma and its relatives,
but many members of the suborder Bodina live in
freshwater (Vickerman, 1976). They have a unique
single mitochondrion which runs the length of the cell
and one or more DNA-rich organelles, kinetoplasts,
associated with this mitochondrion near the flagella.
The best known genus is Bodo, which, like other
bodonids, has two flagella (Fig. 14J). One trails, often
in contact with the substrate, while the other extends
ahead. The trailing flagellum may attach temporarily to
the substrate. In the related, but sessile and colonial,
genus Cephalothamnium (Fig. 14L), the attachment is
permanent.
Diplomonads and cercomonads are relatively
unimportant groups to the freshwater ecologist;
diplomonads are largely parasitic, with a few freeliving forms that may be found in organically enriched
water (e.g., Hexamita, Fig. 14M). There is one freshwater cercomonad, Cercomonas (Fig. 14K), and a common soil flagellate, Heteromita. Cercomonas, like the
diplomonadida, tends to be found in organically enriched environments.
The cryptomonads include many common heterotrophs and autotrophs; mixotrophy has been reported, but is not common. The two flagella are unequal in length and different in appearance (Fig. 7).
The longer one has two rows of mastigonemes, while
the shorter has only one. These flagella arise from a
subapical invagination commonly referred to as a
“gullet,” although it does not appear to be the site
of ingestion in heterotrophic forms. The contractile
vacuole empties into this vestibule. Ejectisomes (see
above) may not be visible to the light microscopist
unless they are discharged, which they sometimes are
after fixation. The pellicle is covered with plates,
although these also are not generally visible.
The dinoflagellates (Fig. 13A – C) are a very large
and unique group; they are probably more important
in marine than in freshwater environments. Certainly,
our knowledge of this group is mostly based on marine
studies. Their unique arrangement of flagella — one
spirals around the cell in a groove (girdle) and the
second is distally directed in another groove (sulcus) —
makes them distinctive. Again, heterotrophy and
mixotrophy are common (Fig. 2). Because dinoflagellates sometimes ingest large prey, their phagotrophic
habits have been more often recognized. A covering of
plates may or may not be present (hence “armored”
and “naked” dinoflagellates). Ingestion in some species
is assisted by an appendage, the peduncle, which
emerges from the sulcus and allows them to subdue
prey many times their size (Gaines and Elbrächter,
1987; Jacobson and Anderson, 1986). The trichocysts
also assist in prey capture in some forms. Very large
prey may be digested by protoplasmic extensions, and
this tendency intergrades with ectoparasitism, which is
common in this group. Marine and brackish species are
noted for the production of toxins.
Chrysophytes also contain both colorless heterotrophs and pigmented mixotrophs. Even among the
mixotrophs, the degree to which different taxa rely on
phagotrophy varies greatly. Chrysophytes are generally
small, and their prey are bacteria. They have two
unequal flagella: one long and directed anteriorly with
rows of mastigonemes; and the other short, smooth,
and directed laterally (Fig. 4D, Fig. 13K – M). They are
naked or covered with fine siliceous scales which
are not always visible with light microscopy; many are
amoeboid. Their carbohydrate storage product, chrysolaminarin, occurs in liquid globules and may be useful
in recognizing members of this group.
Euglenids are generally large flagellates with two
flagella, although in many taxa only one flagellum
emerges from the gullet (Fig. 13D). Several heterotrophic species creep over the substrate with the
second flagellum trailing and hidden beneath the cell
(Fig. 15F – H) as in some bodonids and Cercomonas.
Indeed, molecular genetic studies indicate that the euglenids may be related to the kinetoplastids, and both
may be rather remote from other flagellates (Gunderson et al., 1987). Heterotrophic species may have an
ingestatory apparatus consisting of two rods, allowing
them to swallow relatively large items. Again, the gullet
is not used in ingestion.
3. Protozoa
FIGURE 8
Types of pseudopodia. (A) The cylindrical pseudopodia
of Amoebidae. (B) The cylindrical, granular, eruptive pseudopodia of
Pelomyxidae. (C) The flattened, hyaline, eruptive pseudopodia of
some Entamoebidae. (D) The axopodia of Heliozoa. (E) The filiform
pseudopodia of some Mayorellidae. (F) The broad, flattened
pharopodia of some Mayorellidae. (G) The filopodia of athalamids.
(Modified from Bovee and Jahn, 1973.)
53
around the cell. Locomotion may be achieved by
extending many pseudopodia simultaneously, as in
Amoeba (Fig. 15F), or by moving as a single mass on
a broad front (e.g., Figs. 8F, 15E,P), or as a cylinder
(limax amoebas, Figs. 8A, 15I,K,L). Not only do
pseudopodia have characteristic shapes, but the tail end
or a uroid may be distinctive (Fig. 15J,L), and the cell
surface may be distinctly sculptured, as in Thecamoeba
(Fig. 15D). The dynamics of movement also varies;
some amoebas move continuously and smoothly, while
the cytoplasm of others seems to burst forward intermittently; this motion is called eruptive.
Members of the subclass Testacealobosia are
distinguished by their external coverings, called tests
(Fig. 16A – Q). These are generally vase-shaped, with a
single opening through which pseudopodia emerge.
The test may be proteinaceous, but is often covered
with secreted plates or inorganic particles. Many are
terrestrial, but benthic forms are common and a few
are planktonic. Planktonic species have oil droplets or
gas-filled vacuoles to compensate for the weight of the
test (Meisterfeld, 1991). Others use the gas vacuoles,
which can appear in a matter of minutes, to leave the
substrate and rise to the surface film where pseudopodia are extended at the air-water interface (Ogden,
1991).
Schizopyrenida are small amoebas, many of which
are capable of transforming into flagellates (Fig. 9C).
D. Anatomy of Amoeboid Protozoa
These are the protozoa we generally refer to as
amoebas, in reference to the well-known Amoeba proteus. Locomotion is mostly by pseudopodia. Flagella
are restricted to temporary swimming stages in the
order Schizopyrenida. Rhizopoda and Karyoblastea
mostly have blunt pseudopodia called lobopodia,
where as the Filosea and Granuloreticulosida have
pseudopodia ending in fine points (Fig. 8). Amoebas
range in size from only a few micrometers to 2 mm in
diameter. Although many lack a fixed external morphology, the characteristic morphologies shown by the
various taxa are surprisingly distinctive even if difficult
to quantify (Fig. 15). By using also the number, size,
and structure of organelles and characteristics of tests
(where present), identification is not as difficult for
living specimens as might be imagined.
The morphology of amoebas is plastic. Many
adopt a stellate morphology (Fig. 9) if suspended in
water. Few are truly planktonic, but rather they live on
surfaces or in sediments. Most show an inner granular
cytoplasm and an outer hyaline cytoplasm, or hyaloplasm, with a characteristic thickness and distribution
FIGURE 9 Some amoebas and their “floating forms” on the right.
(A) Ameoba proteus, scale 100 m. (B) Polychaos timidum, scale
20 m. (C) Naegleria gruberi, scale 10 m. (After Page, 1976.)
54
W. D. Taylor and R. W. Sanders
The flagellated form may have two or four flagella. As
amoebas, they are small, uninucleate, and usually
simple in form. The amoeboid form may be a soildweller, transforming into a flagellate on being suspended in water. The best known amoeboflagellate of
this kind is Naegleria. Naegleria gruberi was considered by Page (1976) as possibly the most common
amoeba in freshwater. At one time, Naegleria was best
known as a model system for microtubule assembly,
since it could be induced to assemble its flagellum in
the laboratory. The similar Naegleria fowleri was discovered as a cause of fatal encephalitis in humans in
1958, and the genus is now better known in infamy as
an unwanted resident of air conditioners and swimming pools. Pathogenic strains are able to survive at
37°C, which, fortunately, most free-living amoebas
cannot. Increased abundance of pathogenic N. fowleri
is a dangerous consequence of thermal pollution
(Tyndal et al., 1989). The interesting history of this discovery, as well as other aspects of this group, can be
found in Carter (1968) and Singh (1975).
Filosea and Granuloreticulosida are amoebas with
fine, pointed pseudopodia. Filosea (Figs. 16S – U,W,
19A – K) have single or branched filopodia with fine
points (Fig. 8G), while Granuloreticulosida have a network of branching and anastomosing reticulopodia
(Fig. 17L – R). Most members of these two classes have
tests. The freshwater granuloreticulosids are among the
least studied protozoa, although their marine relatives
are very well known.
Naked amoebas feed by phagocytosis, the smaller
being bacterivores and the larger being predators of algae, other protozoa, and small metazoa. Particulate food
is surrounded by ectoplasm, sometimes engulfed by a
pseudopod, and enveloped in a food vacuole. In contrast
to most other phagotrophic protozoa, there is no fixed
site of ingestion or cytostome. Testacealobosia are
largely algivores, and many feed on filamentous algae,
drawing in filaments through the opening in their test.
Pinocytosis or “cell drinking” can be induced in
Amoeba by dissolved organic substances. Channels appear in the ectoplasm through which substances concentrated on the cell surface are imbibed. Although
Amoeba does not have mucocysts, a mucous coat on
the cell surface appears to be involved (ChapmanAndresen, 1973). The significance of this form of nutrition in the field is unknown, but one must conclude
that Amoeba uses this ability somewhere other than on
microscope slides in undergraduate laboratories!
E. Anatomy of Heliozoa
The Heliozoa are primarily freshwater counterparts of the better known marine Radiolaria (classes
Polycystinea and Phaeodarea) and Acantharia. All
these groups have unique pseudopodia, called axopodia (Fig. 8D). Axopodia are strengthened by a microtubular array called an axoneme or stereoplasm. [The
term axoneme is also used to describe the microtubular
core of cilia and flagella, but this does not imply homology.] The origin and ultrastructure of axonemes is
diverse; indeed, the Heliozoa may well be polyphyletic.
Most Heliozoa lack the skeleton that is so characteristic of Radiolaria and Acantharia, although some are
covered in siliceous or organic scales (Fig. 18H,F), and
some have a perforated shell or capsule (order
Desmothoracida, Fig. 18A).
As diverse as the evolutionary origin of the stereoplasm may be, the axopodia supported by the microtubles are similar in function. Their surface is sticky,
possibly because of extrusomes on the axopodia or
near their bases (Fig. 7). The outside cytoplasmic layer
of the axopods appears to be in constant motion,
streaming out and returning on opposite sides of each
axopod. Food items adhere and are transported with
the flow of cytoplasm until they reach the surface of
the cell body and are engulfed. In this way, they are
functionally analogous to the microvilli of choanoflagellates. Food items may range from picoplankton to
mesozooplankton in different Heliozoa. Large prey
items, such as rotifers and copepods in the case of Actinosphaerium, may be entangled in several axopodia
and engulfed by pseudopods. Several individuals may
participate in the capture of one prey.
Although Heliozoa are frequently planktonic, they
are found primarily on or near the benthos. Some
Heliozoa traverse the bottom with a unique tumbling
motion resulting from controlled changes in the length
of the axopods. Many sessile forms with stalks are
known. In sessile forms, cell division is likely to be unequal, producing a dispersal stage that may be flagellated or amoeboid.
F. Anatomy of Ciliates
Unlike many of the other protozoan groups, the
phylum Ciliophora is certainly monophyletic; despite
their diversity, they share many features. After discussing these common characteristics, we will consider
the important differences among subphyla and classes,
emphasizing organelles involved in locomotion and
feeding.
The presence of cilia is an obvious and distinctive
feature. The cilia may be reduced in number, especially
in sessile forms, or organized into larger compound ciliary organelles, such as cirri. The only large group that
does not always possess cilia is the Suctoria, sessile
predators whose dispersal stages are, however, ciliated.
3. Protozoa
This distinctive group is easily recognized by its feeding
tentacles. The novice should take care not to confuse
small, ciliated animals with ciliates; the size range of
ciliates overlaps that of several metazoan groups, such
as turbellarians, rotifers, and gastrotrichs.
The cortex of Ciliophora is a wonderful example
of the complex cell ultrastructure to be found among
protozoa. We refer the reader to Lynn (1981) and Lynn
and Corliss (1990). This ultrastructure is important in
the definition of higher taxa, while the arrangement of
both somatic cilia and the compound ciliary organelles,
including those associated with the cytostome, are important at all taxonomic levels.
The other distinctive characteristic of ciliates is
their unique nuclear organization or nuclear dualism.
One or more large amitotic, highly polyploid macronuclei are functional in RNA synthesis, while one or more
smaller, diploid micronuclei function as gametic nuclei
during conjugation (see Section III.B).
The ciliates are divisible into ten classes (Small and
Lynn, 1985). Class Karyorelictea is thought primitive
for the group, with numerous macronuclei that are not
highly polyploid. They are largely benthic. The freshwater genus Loxodes (Fig. 23J) is perhaps better known in
its environmental physiology and field ecology than any
other heterotrophic protozoan (reviewed in Goulder,
1980; Finlay 1990). Compound ciliary organelles associated with the cytostome are prominent in the classes
Heterotrichea and Spirotrichea. Large heterotrichs, such
as Stentor and Spirostomum (Fig. 22F,A) are familiar as
teaching material. Spirotrichs are abundant in many
freshwater habitats, from plankton (choreotrichs and
oligotrichs, Fig. 21S – W) to anaerobic benthos (odontostomes and armophorids, Fig. 21J – N,R). Most consume algae and other nanoplankton-sized food particles. Stichotrichs and hypotrichs (Fig. 21A – H,N – Q,
20X – Z) are mostly dorsoventrally flattened crawlers,
whose somatic cilia are all compound cirri.
Like spirotrichs and heterotrichs, Nassophorea
tend to be algivores except for one small and poorly
known group, the microthoracids. The Nassophorea
are named for their basket-like “nasse” or cyrtos supporting the cytopharynx (Fig. 12).
Classes Prostomatea and Litostomatea are largely
predators, often of other ciliates(Fig. 22G-W, 23A-I).
Prostomes generally have apical cytostomes, while
many litostomes have subapical, sometimes slitlike cytostomes. The mouth is encircled by a crown of cilia
from whose bases (kinetosomes) arise the “rhabdos,”
a cylinder of microtubules surrounding, and supporting the cytopharynx. Toxicysts are found in most
species and are used to subdue active prey. Toxicysts
may be found around the cytostome, on a proboscis,
on tentacles, or elsewhere on the body. A number of
55
short, specialized kineties (rows of kinetosomes) are
often found near the anterior. This “brosse” (brush)
probably assists in prey recognition.
Class Phyllopharyngea contains the distinctive Suctoria (Fig. 19), sessile or free-floating predators of other
ciliates. Suctoria are unusual in that most have several
feeding tentacles rather than a single mouth. Prey ciliates stick to these tentacles because of the firing of haptocysts (see extrusomes, above), and their cytoplasm is
withdrawn through the tentacles. Suctoria reproduce
by unequal binary fission (budding), which yields a ciliated dispersal stage or “swarmer.” The other subclass,
the Phyllopharyngia, contains surface-associated algivores (e.g., Chilodonella, Fig. 24T), plus a diverse array of marine epizooic forms.
Colpodea (Fig. 24K,L,N,P – S) are not as frequent
in freshwater environments, most being terrestrial bacterivores. They are more likely to be encountered in
small, temporary waters, and can be readily obtained
by incubating terrestrial vegetation or soil in water. Plagiopylea (Fig. 24M) were formerly placed in the
Colpodea, and they do resemble colpodids in form. Unlike colpodids, they are strict anaerobes.
The Oligohymenophorea are mostly microphagous.
These are named for the compound ciliary organelles
that are found in a buccal cavity surrounding the cytostome. The most common pattern (subclasses Hymenostomatia, Scuticociliatia, Peniculia; Figs. 24C,H,
23L – X, 24A – E,G,I,J) comprises three polykinetids on
the left side of the buccal cavity and an undulating
membrane on the right. The net result is three brushes,
the polykinetids, working against a curved wall, the
undulating membrane, to deliver small particles to the
cytostome. Most are effective bacterivores, although
some are histophagous (consuming tissue of injured or
recently killed animals), parasitic, or algivorous. The
large subclass Peritrichia (Fig. 20A – U) contains sessile
bacterivores in which the buccal cavity is deepened as
an infundibulum, and the polykinetids wind down it to
the cytostome after encircling a prominent peristome.
Somatic ciliature is absent in most species. Many are attached to the substrate by a contractile stalk, as in the
common Vorticella. Many are colonial, and a few secondarily free-swimming.
G. Environmental Physiology
1. Factors Affecting the Distribution of Protozoa
Protozoa are found over an extremely broad range
of environmental conditions, but individual species,
especially those adapted to unusual circumstances, may
have relatively narrow limits to their distribution.
Several microbial communities with characteristic
56
W. D. Taylor and R. W. Sanders
fauna have been described. Biological, physical, and
chemical factors must interact to govern the distribution of these assemblages in a manner that defies simple
description. Nonetheless, several physical – chemical
factors are important to the growth and/or distribution
of protozoa. Among these temperature and oxygen are
probably the most important and best known. Light is
obviously important to autotrophic forms, and the negative influence of UV light on protozoa is probably an
important factor that has received too little attention
(Sommaruga et al., 1996).
An extensive and still rapidly expanding literature
concerns the importance of environmental contaminants. The distribution of protozoa with respect to organic pollution and its associated environmental conditions has been used as part of a system for classifying
the state of aquatic habitats according to their fauna
(Sládecek, 1973). Foissner (1988) and Foissner and
Berger (1996) have clarified the taxonomy of ciliates
used as indicators and provided a summary of the
“saprobity values” of ciliates. Similarly, Bick (1972)
provided a guide to ciliates useful as biological indicators in freshwater, along with their ecology and distribution with respect to several parameters.
A list of species occurrences versus environmental
parameters was assembled by Noland (1925). His conclusion that “the wide range of tolerance that most ciliates show toward the physico-chemical factors studied,
together with the evident correlation of several of these
factors with the food habits of the species, suggests that
the nature and amount of available food has more to do
with distribution of the freshwater ciliates than any
other one factor” still appears to be warranted. The
physical nature of the substrate, including the biological
substrate of periphyton, is also important to the community of surface-oriented protozoans that develops on
it (Picken, 1937; Ricci, 1989). Indeed, the relative importance of biotic components of the environment to
the distribution of species versus the structural and
chemical – physical components, is difficult to ascertain
from survey data.
2. Temperature
As in all organisms, metabolic processes and
growth in protozoa are dependent on temperature. Because of the difficulty of measuring growth rates in situ
for protozoa, several workers have investigated the
relationship of temperature and growth rate in the laboratory to facilitate the extrapolation of laboratory
data on growth rate to field situations (Finlay, 1977;
Baldock et al., 1980). Generally, Q10 values for protozoa are about 2.0, as they are for many biological
processes, but important deviations occur (Baldock and
Berger, 1984; Müller and Geller, 1993).
Active protozoa are found in freshwater environments at temperatures between 0 and 50°C. For example, protozoa contribute to the plankton community of
almost permanently ice-covered lakes in Antarctica
(Laybourn-Parry et al., 1995). Some protozoa can be
cultured in the laboratory at or near 0°C, but it is
doubtful whether all protozoa can function at such low
temperatures, or whether those that can perform any
better than one would expect from extrapolating the
metabolic rate versus temperature curve for species collected from more moderate temperatures to 0°C. Lee
and Fenchel (1972) found that Euplotes collected from
Antarctic, temperate, and subtropical waters had similar growth – temperature relationships; but the Antarctic strains did not multiply above about 12°C, while
the subtropical strain multiplied even above 35°C, but
could not reproduce below 6°C.
A similar situation is to be expected with respect to
high temperatures; most species continue to increase their
rate of cell division as temperature increases to 25°C or
more and tolerate at least 30°C. But there are numerous
records of species from hot environments, up to 50°C
(Noland and Goldjics, 1967). Tolerance to extremes of
temperature and some other variables have been induced
by gradual acclimation (see Fauré-Fremiet, 1967).
3. Oxygen
Protozoa are sufficiently small that they do not
need organelles to increase the surface area for respiration. Aerobic forms appear to function at extremely
low oxygen tensions and, thereby, avoid competition
and predation from larger animals. It is in these circumstances, especially when they are generated by high densities of bacteria in response to organic enrichment, that
protozoa are most abundant. Aerobic protozoa that
are typically found under such conditions are called
microaerophilic. The succession of species following organic enrichments and the resulting changes in oxygen
tension were a focus for some of the earliest research in
protozoan ecology (Noland and Goldjics, 1967).
A great deal of research has been done on the
autecology of Loxodes, a microaerophilic ciliate living
in the benthos and plankton of hypereutrophic waters
where it consumes nanoplanktonic algae (Fenchel and
Finlay, 1991). One of the most surprising outcomes of
this research is the discovery that Loxodes can use
nitrate rather than oxygen as an electron acceptor. It
undergoes diel vertical migrations between the anoxic
hypolimnion and oxygenated epilimnion, but cannot
survive in either if constrained. It aggregates at low O2
concentrations guided, in part, by unique organelles
(Müller’s vesicles) now known to be responsible for
geotaxis. The direction of the geotactic response is
determined by the oxygen tension, which is probably
3. Protozoa
detected by cytochromes, and by light, which is
detected by pigment granules (pigmentocysts) in the
pellicle. Although other ciliates have been intensively
studied in the laboratory, the work on Loxodes is unusual in the degree to which laboratory studies have
been guided by field observations. This suggests that
autecological work on other species could be very
rewarding.
Some microaerophilic ciliates contain endosymbiotic green algae (see Section III). Despite their photosynthetic symbionts, they aggregate at depths where O2
and light are low (Finlay et al., 1987). At very low O2
tensions, they move into dim light suggesting a role for
the symbionts not only as producers of carbon for their
hosts but as a means for maintaining a low intracellular level of oxygen.
Many free-living protozoa are tolerant of anaerobic conditions (Fenchel and Finlay, 1991). Among the
ciliates, this diverse assemblage of species have been
referred to as “sapropelobionts” (e.g., Jankowski,
1964a,b), although some of the species embraced by
this term are microaerophilic rather than anaerobic.
Fenchel (1969) suggested that the term “sulfide ciliates” is more appropriate for the truly anaerobic ciliates of reducing sediments. These are intolerant of oxygen and lack mitochondria and cytochrome oxidase.
Instead of mitochondria, some anaerobic protozoa
have hydrogenosomes, organelles that resemble mitochondria, but reduce pyruvate and generate hydrogen.
Epi- and endozooic bacteria are commonly associated
with anaerobic protozoa, and methanogenic endosymbionts, in particular, may be closely associated with the
hydrogenosomes (Fenchel and Finlay, 1991). Ecological
studies of freshwater species have appeared in recent
years (Wagener et al., 1990, Massana and Pedrós-Alió,
1994; Guhl et al., 1996).
Other protozoa which can probably exist under
anaerobic conditions include the giant amoeba
Pelomyxa palustris, which has been said to consume at
least some oxygen (Bovee and Jahn, 1973) although it
has no mitochondria (Daniels, 1973). Oxygen consumption in some anaerobes may be a protective mechanism (Fenchel and Finlay, 1991). The diplomonads,
which have two free-living genera (Hexamita and Trepomonas) also lack mitichondria. Flagellated protozoa
in the anaerobic hypolimnia of eutrophic lakes may
reach abundances as great or greater than those in the
surface waters (Bennett et al., 1990). Many true anaerobes among the protozoa belong to taxa that include
gut-dwelling endosymbionts.
4. Environmental Contaminants
The relative abundance and diversity of protozoa
has also been proposed as a useful indicator of organic
57
and toxic pollution (Cairns et al., 1972), and the
response of protozoa to contaminants has been reviewed in that light (Cairns, 1974). Others have advocated the use of protozoa in single-species bioassays
(Nilsson, 1989). Relatively quick respiration bioassays
may reveal both decreases and transient increases in respiration (Slabbert and Morgan, 1982). Because tests
with protozoa are relatively rapid, they are particularly
useful for studies of quantitative structure – activity
relationships (QSAR) where the toxicity of many compounds must be compared. The response of Tetrahymena to a wide variety of compounds is similar to that
of a cyprinid, the fathead minnow (Bearden and Schultz,
1997). Protozoan predators and prey have been used to
explore complex effects of contaminants on communities (Doucet and Maly, 1990, Fernandez-Leborans and
Novillo-Vajos, 1993).
Among environmental contaminants, the response
of protozoa to oil and other hydrocarbons is relatively
well known. They are generally resistant to hydrocarbons (Rogerson and Berger, 1981); the ciliate Colpidium colpoda accumulated hydrocarbons in intracellular
inclusions, up to about 20% of the cell volume. The
number of mucocysts increased, implicating these organelles as having a protective function (see Section
II.B.4). Scott et al. (1984) found that freshwater ponds
treated with oil alone or oil and oil-dispersant mixtures
had numbers and biomass of protozoa similar to those
in control ponds, although some taxa were reduced
while others increased.
The toxicity of various hydrocarbons to Tetrahymena was similar when differences in solubility and
partitioning between the water and ciliates were considered, indicating a similar mode of action once they
were in the cell (Rogerson et al., 1983). Biomagnification of chlorinated hydrocarbons may be substantial.
For the pesticide Mirex, the concentration in Tetrahymena was 193 times that present in the medium, and
82% of the available contaminant was in the cells
(Cooley et al., 1972). For Arochlor™ 1254, the biomagnification was 60 -fold. A wide variety of insecticides have been shown to cause dose-dependent inhibition of growth and biomagnification in Tetrahymena
(Dhanaraj et al., 1989; Lal et al., 1987). Some were
toxic at higher concentrations.
The toxicity of heavy metals to protozoa also has
been explored in some detail (Ruthven and Cairns,
1973; Doucet and Maly, 1990; Fernandez-Leborans
and Novillo-Vajos, 1993). Intraspecific variation in
heavy metal tolerance appears to be related to environmental variation in contamination (Nyberg and Bishop,
1983). To some degree, tolerance to metals is specific,
but is also generally greater in outbreeding species
(Nyberg, 1974; see Section III.B for discussion of
58
W. D. Taylor and R. W. Sanders
breeding systems). Free-ion concentration, rather than
the total amount of metal, appears to determine toxicity (Stoecker et al., 1986).
5. Energetics of Protozoa
Because protozoa are fast-growing and easily cultivated in the laboratory, their energetics have been studied extensively. Ingestion, growth, respiration, egestion,
and other parameters have been measured or estimated, and quotients cataloged. We deal with some of
these parameters elsewhere; for example, maximum
growth rates are discussed in Section II.B, some field estimates of growth rate are mentioned in Section III.B
and egestion in Section III.C.10). Laybourn-Parry
(1984) summarized much of the existing information
on protozoan energetics, and Calow (1977) integrated
it into his wider review of animal energetics.
Fenchel’s (1974) classic paper drew attention to the
relationship between body size and rate of population
increase for organisms, and the more rapid growth rate
of the smaller as compared to the larger ones. His
analysis also suggested a cost associated with multicellularity; both net growth efficiency (growth/assimilation) and growth are generally larger for protozoa than
for metazoa of similar size. Calow’s summary of conversion efficiencies confirms that protozoa are generally
efficient converters of energy, with many species showing growth efficiencies of 50% or more. However, efficiencies for some taxa are much lower (e.g., Scott,
1985, and references therein) and it is difficult to generalize. Given the high growth rates of protozoa, we expect that field populations of protozoa will be found to
have high production-to-biomass (P/B) ratios relative
to metazoa, on the order of one per day. However, few
field estimates of growth rate are available. Field estimates of growth rates include Schönborn (1977) and
Taylor (1983b) for benthic populations, and Taylor
and Johannsson (1991) and Carrick et al. (1992) for
planktonic populations. It appears that planktonic ciliates undergo periods of consumer and resource control,
and these may be different for different species (Havens
and Beaver, 1997; Wang and Heath, 1997).
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
The concept of species within the protozoa has
always been problematic. The definition of species as
interbreeding or potentially interbreeding populations
is irrelevant to the many asexual forms. On the other
hand, studies of the genetics of what were once believed to be species, such as Paramecium aurelia,
Tetrahymena pyriformis, and Euplotes patella, have revealed groups of interfertile mating types (Corliss and
Daggett, 1983). These groups of mating types are
called syngens by protozoologists, but are essentially
sibling species. New species names have been created to
recognize them in some cases (e.g., P. monaurelia, P.
biaurelia, and so on). Careful morphological analysis
has uncovered differences among some of these species,
but in practice, they are not morphologically separable.
They must be identified by mating reactions with
known lines, or by using electrophoretic differences in
some cytoplasmic enzymes. At the same time, the phenotypic plasticity of protozoa has also led to the description of invalid species names (Gates, 1978). Finlay
et al. (1996) provided a carefully reasoned analysis of
this problem, and concluded that “the biological
species concept is neither appropriate nor practicable”
for ciliated protozoa. They concluded that we must instead be pragmatic and consider morphological species.
The same would be more true of phyla where sex is less
common or absent. It remains that some protozoan
groups have as many species as the more familiar
animal taxa. For example, ciliates comprise some 7000
species (Corliss, 1979), with about 3000 known freeliving species (Finlay et al., 1996), of the tens of thousands of protozoan species.
At this point it should be admitted that the taxonomy of free-living protozoa, at least as practiced by
ecologists, is in a primitive state. A textbook on protozoa will advise the student to collect protozoa at the
margin of a pond or in some other small and protected
waters rich in organic material. Collections from these
environments are likely to yield rich assortment of
large protozoa, readily identifiable at least to genus under the dissecting or compound phase microscope. The
working freshwater ecologist will more likely be interested in lake plankton, littoral or profundal benthos, or
stream aufwuchs. Samples from these situations will
more likely yield minute forms too small to be seen under a dissecting microscope, and too few to be examined under a compound microscope without concentration. They may not be figured in the textbooks. Many
will be undescribed species, or species descriptions will
be available only in specialist journals, not to be found
in most libraries, or so outdated as to be of dubious
value. Largely because of these problems, most ecologists who include protozoa in their studies of aquatic
habitats do not identify protozoa, even if they do count
them and measure them for biomass estimates. Techniques of sample preservation, concentration, and enumeration (see Section IV), if they exist for a particular
biotope, may not allow for identification. Molecular
techniques such as species-specific oligonucleotide
probes have been developed and may prove to be
3. Protozoa
useful tools for identification of particular protozoan
groups in natural samples (Lim, 1996).
The distribution of most free-living protozoa is
probably broad. Certainly, some morphological species
are worldwide in distribution and are essentially ubiquitous. Experiments using enriched and incubated field
samples led Fenchel et al. (1997) to conclude that although local diversity of protozoa may be high, global
diversity may not be; morphological species are usually
present in appropriate habitats, even if not detected by
initial sampling. Protozoa quickly colonize new environments, as was shown by Maguire’s (1977) study of
new volcanic islands. On the other hand, sibling species
of the Paramecium aurelia complex have limited distributions on a world wide scale (Sonneborn, 1975) even
though the species complex is ubiquitous.
B. Reproduction and Life History
Most protozoa reproduce by equal binary fission,
and therefore their life histories are simple. Multiple fission is more common among endosymbionts but does
also occur in free-living forms. Among free-living protozoa, multiple fission appears to be related to intense
temporal heterogeneity in the environment. For example, it is found in some terrestrial forms which are active
only for brief periods when their environment is moist,
or in histophages whose food resources are very patchy
and ephemeral (Fig. 10). Therefore, some analogy can
be drawn between the significance of multiple fission in
certain free-living protozoa and some endosymbionts
using the common theme of dispersal (Taylor, 1981).
A phenomenon perhaps related to multiple fission,
in that it is related to dispersal, is unequal fission or
budding. Binary fission is only approximately equal,
and differences in the partitioning of cytoplasm occur
and are adjusted in the next cell cycle. But a consistent
asymmetry of daughter cells is widely distributed
among sessile forms; the larger daughter cell remains
attached, while the smaller is motile. This pattern undoubtedly has evolved independently a number of
times. It is perhaps a reflection of the probability of survival of the two daughter cells not being equal, and resources being partitioned according to the odds. The
smaller daughter cell is usually a propagule, capable of
establishing itself in a new location. In some cases,
(e.g., Zoothamnium, Fig. 20H,I), it may function as a
gamete.
Generation times are highly variable among protozoa, but the maximum rates of population increase
attainable when food is not a constraint generally increase with temperature and decrease with body size
(see Section II.5). Multiple regressions of growth rate
on cell size and temperature are sometimes used to
59
estimate production despite shortcomings (Müller and
Geller, 1993). However, even for protozoa of the same
size and growing at the same temperature, maximum
rates of cell division vary widely among taxa. Some of
this variation is correlated with genome size and may
reflect a causative relationship. Among most eukaryotic
cells with diploid, mitotic nuclei, there is a positive relationship between DNA content and cell size, and a
negative relationship between DNA content and division rate. It appears that, among such cells, a large cell
must have a proportionately large DNA content and
nucleus, and a slow division rate. This relationship
holds among most plant and animal cells and among
most protozoa. However, it has also been established in
plants that polyploidy allows a large DNA content and
large cell size, but without a concomitant decrease in
division rate. Division rate is apparently related to the
size of the genome, not the number of copies.
The diverse nuclear arrangements of protozoa can
be viewed as various solutions to the genome size/cell
FIGURE 10
Life cycle of the histophagous ciliate Ophryoglena.
Change from the theront (hunting morphology) (A) to the trophont
(feeding) morphology (B,C) occurs when food is found. The prototomont stage (D) seeks a suitable place to settle, and leads to the cyst
or tomont stage (E,H). The tomont usually produces 4–8 new
theronts, or may produce a much larger number of microtheronts
(F,G). The function of microtheronts is unknown. Theronts may encyst if they do not find food, and re-emerge as small secondary
theronts. (Modified from Canella and Rocchi-Canella, 1976.)
60
W. D. Taylor and R. W. Sanders
size/fission rate constraint (Cavalier-Smith, 1980).
Multinucleate cells, nuclear dualism, amitotic nuclei,
and colony formation may represent ways to get larger
without slowing growth. For example, in ciliates, cell
size is correlated with macronuclear DNA content, but
cells divide as frequently as one would expect from the
size of their micronucleus (Shuter et al., 1983). In this
context, both multinuclearity and multicellularity can
be seen as means of avoiding the constraints imposed
by the basic eukaryotic plan.
Encystment is a feature of many protozoan life
cycles, especially those that live in environments that
dry out (Corliss and Esser, 1974). Many protozoa also
produce cysts at times when environmental conditions
are unsuitable for other reasons, such as lack of food.
Encystment under these conditions can be considered
as a means of coping with temporal heterogeneity. Still
other protozoa incorporate encystment into the life
cycle following feeding (the “digestive cyst”) or during
cell division (the “reproductive cyst”). Digestive cysts
are common in ciliates that rapidly consume large
amounts of food, possibly after a long period of searching, e.g., histophages and predators. Some species enter
a cyst after feeding and emerge after cell division, combining these functions. The encysted stage may reduce
energy expenditure and/or the risk of predation when
the acquisition of food is either not possible or unnecessary. Nonencysting protozoa may persist days or
weeks in the absence of food; this survival time is not
closely related to cell size, is density-dependent, and
probably reflects differences in energy costs associated
with locomotory behavior (Jackson and Berger, 1985).
Survival time was found to be negatively correlated
with intrinsic rate of population increase (see Section
III.C.2) among eleven species, but there were exceptions to this trend. Crawford and Stoecker (1996) examined survival and respiration during starvation in a
marine mixotrophic ciliate Strombidum capitatum.
There was no decrease in dark respiration with starvation, and the starving population comprised both
larger cells with low specific respiration rates and small
cells with high specific respiration rates. Survival time
was longest for the large cells.
Sex is widespread, but far from universal among
protozoa. For example, it is common among the
Ciliophora, Chlorophyta, Chrysophyta, and Granureticulosida, limited among Dinoflagellata and Heliozoa, and absent in most if not all Zoomastigina and
Rhizopoda. Grell (1973) and Margulis and Sagan
(1986) discussed the occurrence and variety of sexual
phenomena among the protists. Grell organized sexual
phenomena into three basic types: (1) gametogamy, in
which gametes fuse as free-swimming cells; (2) autogamy, where gametes or gametic nuclei of the same
individual fuse; and (3) gamontogamy, where mates
(gamonts) unite to exchange gametes or gametic nuclei.
Gametogamy is found in flagellates where, typically,
isogametes are produced by multiple fission. However,
anisogamy also occurs. Autogamy is present in Heliozoa and some Ciliophora. Gamontogamy is the usual
form of sex in Ciliophora.
As in most small metazoa, sexuality interrupts
asexual reproduction, and is associated with environmental change. It should be emphasized that sexuality
is not necessarily associated with reproduction in protozoa and is best understood as a separate phenomenon
(Margulis and Sagan, 1986). Indeed, sex usually marks
the end of the existence of an individual protist; it becomes the gamete or gametes, and its genome ceases to
exist. The restricted occurrence of sex among protozoa,
and the separation of sex from reproduction in many
taxa, make protozoa a promising group for further
study concerning the evolution and significance of sex.
The relationship between sex and asexual reproduction has been carefully examined in some Ciliophora, especially Paramecium (see Wichterman, 1986,
for a review of this topic). Exchange of haploid nuclei
between two conjugants (gamontogamy) leads to two
individuals with new and different genotypes. Each is
the founding member of a new clone, expanding by
asexual binary fission. Individuals of a new clone will
undergo a period of “sexual immaturity” when sex
does not occur. The duration of this immature period
varies among species; species with short immature periods are considered to be “inbreeders,” as mating with
close relatives is favored, whereas species with long
immature periods are considered to be “outbreeders.”
The immature period is followed by a period of
sexual maturity when conjugation will occur under the
following circumstances: (1) a partner of a complementary mating type is available; (2) both are at the right
stage of the cell cycle; and (3) both are slightly starved.
If conjugation does not occur before the end of the period of maturity, a period of sexual senescence follows.
The rate of binary fission declines, cells are less likely
to survive conjugation if a mate is found, and autogamy (self-fertilization) may occur. Although autogamy
may prolong vigor, the clones ultimately die out. An
analogy can be drawn between the life of a clone of
ciliates and the development of a metazoan from a
zygote — a large population of cells develops asexually
from the single cell resulting from sexual fusion, but
ultimately the size and life span of that population are
limited by senescence which must be circumvented by
another sexual episode.
As some ciliates do not undergo conjugation and
others vary greatly in the frequency necessary to prevent senescence (a strain of Tetrahymena without a
3. Protozoa
micronucleus and, therefore, not capable of sex was
isolated in 1923 and is still going strong), one is
tempted to speculate on the function of both sex and
senescence. Is sex required to postpone senescence by
replacing irreparably damaged DNA by recombination? Or, is senescence a mechanism to eliminate
micronuclear or gametic genes that have not been expressed for many generations? Or, is the need for sex a
function of the unique amitosis of the ciliate macronucleus? The finite life span of animal cells in tissue
culture versus the immortality of cultures of cancer
cells begs similar questions. That autogamy in ciliates is
a stop-gap measure to postpone senescence contrasts
with sex in the heliozoan Actinophrys sol, where fusion
of the products of meiosis within the same cell is the
normal pattern. Bell (1988) reviewed this fascinating
subject and concluded that senescence of both sexual
and asexual protists can be viewed as a common phenomenon, explicable in terms of the genetic deterioration of finite populations.
Although sex is a necessary event in the life cycle
of some ciliates, it is a dangerous undertaking; genetic
load is high in many ciliates, as is the mortality following conjugation. As noted previously, some species
avoid close matings by having a long immaturity period. As if conjugation was not dangerous enough,
prokaryotic endosymbionts render some ciliates “matekillers”; their partners in conjugation pass on their
genes but are lethally affected (see Section III.C.4).
C. Ecological Interactions
1. Intraspecific Interactions among Protozoa
Protozoa have been used extensively in the study of
competition and predator – prey interactions. Their obvious advantages for this type of study include small
size, short generation time, and, for some species, ease
of maintenance in the laboratory. These advantages do
not carry over to field experiments, and little work has
been done in situ.
Intraspecific competition in protozoa is evident in
the dynamics of batch cultures, where a few individuals
are introduced into a large, but finite environment. If
inoculated into batch cultures as starved individuals,
some ciliates will respond to this situation by growing
rapidly and increasing in size past the biovolume at
which the cell might be expected to divide. This has
been interpreted as a strategy to sequester the maximum amount of food should the supply become exhausted. However, if food is still available, these cells
enter into a phase of exponential growth where they
divide at a similar size in each cycle. When the food
supply is depleted, some individuals will continue to
61
divide at the expense of cell size. Again, this can be
interpreted as an adaptation for an r-selected or colonizing species to produce as many propagules as possible (Taylor, 1981; Fenchel, 1987).
Curds and Cockburn (1971) found that, in order
to model the population dynamics of Tetrahymena in
batch cultures, it was necessary to include interference
among individuals in their equations. This was not necessary for continuous cultures. The exhaustion of food
in some species induces cannibalism, sometimes with
abrupt changes in morphology for the cannibals (see
Giese, 1973, for a thorough discussion of this phenomonen in the ciliate Blepharisma). Another form of
interference is the “killer” phenomenon. Some protozoa harbor prokaryotic endosymbionts that render
them lethal to conspecifics (mate-killers were mentioned above).
2. Interspecific Competition among Protozoa
Interspecific competition in laboratory cultures of
ciliates has been studied by Gause and numerous authors since. Gause (1934) provided clear evidence that
environmental factors could influence the outcome of
competition.
The Lotka – Volterra equations predict that the
winner of exploitative competition for resources in stable environments should be the species with the greater
K or carrying capacity, that is, the more efficient user
of the resource. However, K is usually measured as
numbers, not biomass, so smaller species will tend to
have a higher K. In his studies of competitive interactions of bacterivorous ciliates, Luckinbill (1979) confirmed that K did not predict the outcome of competition. But, in accordance with part of the r/K selection
hypothesis, the intrinsic rate of increase, r, was negatively related to competitive ability. [The r/K selection
hypothesis holds that r, which measures fitness in density-independent or stochastic environments, is negatively correlated among species with K, which measures
fitness in stable environments.] Thus the theorem seems
to hold for ciliates, except that K, being highly correlated to body size, does not measure competitive ability
in stable environments. As noted above, r is also correlated with body size.
To factor out this correlation, Taylor (1978a,b)
expressed intrinsic rate of increase as r/re, where re is
the expected r for a ciliate based on a regression of r on
body volume. Therefore, r/re is a measure of how
fast-growing or r-selected a ciliate is for its body size.
This measure was found to be negatively correlated
with frequency of occurrence in the field, and positively
correlated with macronuclear ploidy (Taylor and
Shuter, 1981). This research suggests that protozoan
communities contain fast-growing but ephemeral
62
W. D. Taylor and R. W. Sanders
opportunists, as well as more conservative, specialist
species that have more modest growth rates, but stable
populations. A similar picture emerged from a comparison of the population dynamics of a relatively fast and
a relatively slow-growing species in the laboratory
(Luckinbill and Fenton, 1978).
Field studies of exploitative competition in protozoa are few, as are field measurements of growth rates
for comparison to laboratory estimates under conditions of excess food. Accordingly, we know little about
the relative importance of density-dependence or independence. Work on planktonic protists suggests that
field populations may experience periods of rapid
growth and periods of little or no growth (Taylor and
Johannsson, 1991; Carrick et al., 1992).
Maguire (1963a,b) found strong evidence for the
exclusion of colonizing species in the genus Colpoda by
Paramecium and associated species. Hairston and
Kellerman (1965) and Hairston (1967) found that one
member of the Paramecium aurelia species complex
dominated sympatric sibling species in competition experiments and predominated in field collections. The
field population they studied appeared to be foodlimited. In contrast, Taylor (1983b) found that sessile
ciliates on an artificial substrate in a stream had high
growth and mortality rates, implying little competition.
The extensive studies on colonization by Cairns and
co-workers (reviewed in Cairns and Yongue, 1977)
provide indirect evidence for species interactions, including competition.
There is also competition between protozoa and
metazoa consuming similar resources. Potential metazoan competitors include some rotifers and crustaceans,
although it is difficult in some cases to distinguish
between the effects of competition and predation
(Wickham and Gilbert, 1993). Likewise, the presence of
predatory protozoa may affect the outcome of competition between other protozoan species (Lawler, 1993).
3. Predatory Interactions among Protozoa
Many protozoa are predators of other protozoa.
For example, Chardez (1985) listed a diverse assortment
of protozoa consuming testaceans. Protozoa have been
used extensively in laboratory studies of predator – prey
interactions, including such topics as selectivity, stability
of predator – prey interactions, and the behavioral and
energetic determinants of functional and numerical response. Much of this research has been done with
Didinium nasutum. Hewett (1980a,b) and Berger
(1979) showed that Didinium changes its body size,
feeding behavior, and feeding and growth rate when fed
different prey and may encounter some difficulty
handling large prey after adjusting to small ones. The
extrusomes which enable Didinium and other predatory
ciliates, dinoflagellates, and Heliozoa to handle their
prey were mentioned in Section II. Although extrusomes
are important weapons of many other ciliates, especially
in the Litostomatea and Suctoria, other predators are
simply swallowers, including the giant cannibal forms
of Blepharisma and Tetrahymena, large ciliates such as
Bursaria, and some other predators such as Peranema
among the Euglenida and the large amoebas.
Interest in the “microbial loop” (the route by
which picoplankton production reaches larger consumers) has focused attention on the contribution of
protozoa, as consumers of picoplankton, to production
at higher trophic levels. A microbial loop leading to
fish via crustacean zooplankton is likely to be inefficient because of the number of steps involved, and even
more inefficient if predatory protozoa process some of
that production. Nevertheless, protozoa are ingested by
many metazoa and often appear to be a high-quality
food (Sanders and Wickham, 1993).
Relatively little is known about the quantitative
significance of predatory protozoa in planktonic food
webs. In some instances ciliates appear to be the major
predators of heterotrophic nanoflagellates (Weisse
et al., 1990), but the relative predation impact on flagellates by ciliates versus metazoa appears to vary seasonally (Weisse, 1991; Sanders et al., 1994). Known
predators of other protozoa, such as Dileptus and Actinosphaerium, are commonly observed in the plankton.
The prey of many abundant litostomes is not known,
but they are likely to feed on other protozoa. Use of
fluorescently labeled prey has identified small ciliates,
including Strobilidium, Halteria, and tintinnids, as
predators of heterotrophic nanoflagellates in lakes
(Cleven, 1996). In some estuarine systems, dinoflagellates may be important predators of ciliates (Bockstahler and Coats, 1993) but this is not documented in
freshwaters. Presence of predatory ciliates, amoebas,
and turbellaria elicits morphological changes in the ciliate Euplotes that reduce predation (Kusch, 1993). Predation within the protozoan microplankton is still an
unknown loss in the transfer of energy to higher
trophic levels and therefore bears on the debate concerning the importance of the microbial loop.
4. Symbionts of Protozoa
Symbionts of protozoa include both ecto- and endosymbiotic prokaryotes and eukaryotes, including a
wide variety of parasites (Curds, 1977; Jeon, 1983; Lee
and Corliss, 1985). The most obvious eukaryote
symbionts of freshwater protozoa are algae of the
genus Chlorella, often called zoochlorellae. These are
most widely studied in the ciliate Paramecium bursaria,
but are also known from a wide variety of other ciliates
as well as flagellates, amoebas, and many metazoa. In
3. Protozoa
63
some cases, the symbiosis appears to benefit the protozoan because of the photosynthates released by the
algae; P. bursaria, for example, probably receives
maltose from its symbionts and can grow better at low
food concentrations when they are present.
Although some alga-bearing protozoa display positive phototaxis and aggregate at the water surface
(Berninger et al., 1986), others aggregate deeper, in dim
light. Some of these microaerophilic forms are negatively phototactic, unless the medium is anaerobic
(Finlay et al., 1987), and zoochlorellae may provide
low internal oxygen tensions in anaerobic external environments. Therefore, there may be two functional
types of protozoan – algal symbioses: one in which the
primary benefit for the host is to receive photosynthate,
characterized by positive phototaxis, and one in which
the primary benefit for the host is oxygen production,
characterized by oxygen-dependent phototaxis.
Some choreotrichs (Strombidum spp.) harbor isolated chloroplasts as symbionts (Rogerson et al., 1989;
Stoecker et al., 1989). The chloroplasts may be from
more than one species of alga. In some cases, whole
cells may be retained as symbionts. The host ciliate
becomes functionally mixotrophic, able to photosynthetically fix significant amounts of carbon. The
haptorid ciliate Perispira ovum ingests the flagellate
Euglena, incorporating its chloroplasts, mitochondria
and paramylon bodies (Johnson et al., 1995). It probably uses oxygen produced by the chloroplasts to live in
low-oxygen environments.
Prokaryotic endosymbionts are widely distributed
in protozoa. For example, several species of anaerobic
protozoa have endosymbiotic methanogenic bacteria
(Section II.H.3). Some endosymbionts are infectious
parasites which may kill their hosts, while some are
necessary for the survival of their hosts. In some cases,
the function of the symbionts is unknown. Perhaps the
most interesting endosymbionts from an ecological
point of view are the intensively studied “killer” particles of ciliates. Some transform their hosts into
“killers” that cause the death of conspecific “sensitives” simply by their proximity. Others transform
their hosts into “mate-killers,” whose partners in conjugation are lethally affected. The significance of these
endosymbionts to field populations of Paramecium
has been investigated by Landis (1981). He concluded
that the infection must be maintained by natural selection, because killers rarely have a chance to mate and
transmit their infection to new hosts. Mating and cell
division are the only known means of transmission.
to be rather nonspecific, involving species that can also
be found attached to, or crawling on, nonliving
substrates as part of the aufwuchs community. Other
associations (e.g., peritrichs of the order Mobilida that
are epizooic on fish) are perhaps better left to the parasitologists. However, some cases deserve brief mention
here because they are common and likely to be noticed.
One common association is between planktonic,
colonial cyanobacteria and peritrich ciliates (Pratt and
Rosen, 1983; Kerr, 1983). Peritrichs are frequently so
abundant as to rival the biomass of their host colony,
and they likely affect its nutrient environment and
motility. Smaller peritrichs, choanoflagellates, and bicoecids are common epizooics on diatoms and filamentous cyanobacteria (Fig. 11A).
Peritrichs and Suctoria colonize the exoskeleton of
planktonic crustacea. Kankaala and Eloranta (1987)
5. Epizooic and Epiphytic Protozoa
FIGURE 11 (A) Choanoflagellates epiphytic on filamentous
cyanobacteria from Georgian Bay, Lake Huron. Scale bar 10 m.
(B) Tokophrya quadripartita (Suctoria) epizooic on Limnocalanus
from Lake Michigan. (From Evans et al., 1979.)
Many freshwater protozoa have developed epizooic
associations with animals. Some associations are likely
64
W. D. Taylor and R. W. Sanders
FIGURE 12 The cyrtos of Nassula, and how it is used to ingest filamentous cyanobacteria. (From Tucker, 1968.)
found that the clearance rates of Vorticella epizooic on
Daphnia can rival those of their host, suggesting that
competition for food with Vorticella could be significant to the Daphnia. Epizooic suctoria (Tokophrya
spp., Fig. 12B) on mysid and calanoid Crustacea in the
Great Lakes may be heavy enough to impair their hosts
(Evans et al., 1981) and may also compete with them
for food. Henebry and Ridgeway (1980) suggested that
epizooic protozoa on Crustacea may be useful pollution indicators.
Benthic invertebrates may also be infested with
peritrichs (see Baldock, 1986, for a brief summary).
The biomass of these may be a significant portion of
the protozoan biomass in streams.
6. Protozoa as Bacterivores
Protozoa are most widely known as bacterivores.
Bacterivorous protozoa are abundant in activatedsludge sewage treatment plants, and, indeed, are neces-
sary for their proper functioning (Curds and Cockburn,
1970). At the opposite extreme, there is considerable
evidence that in plankton communities, especially oligotrophic ones, heterotrophic flagellates are the dominant bacterivores and that much of the carbon flow
passes through them (e.g., Sanders et al., 1989; Nagata,
1988; Carrias et al., 1996). Although ciliates may be
abundant bacterivores in enriched environments (hence,
the old name “infusorians” or animals from organic infusions), they are thought to be quantitatively unimportant in many planktonic ones. However, in some planktonic environments, especially those where cladoceran
crustacea can seasonally reduce flagellate abundances
significantly, ciliates may become the dominant bacterivores (Simek and Straskrabová, 1992). Heterotrophic
flagellates, and to a lesser degree ciliates, also may be
important bacterivores in the water column of rivers
(Carlough and Meyers, 1991), although bacterivory by
protists in this environment has barely been addressed.
3. Protozoa
The relative importance of flagellates and ciliates as
bacterivores in different habitats and during different
seasons is, therefore, an open question. The question
should probably be phrased in terms of the size distribution of grazers of picoplankton, and the answer probably involves species interactions and food web structure
as well as trophic status.
The small Heliozoa are an overlooked group of
largely planktonic bacterivores, and small amoebas are
bacterivorous also. Mixotrophic flagellates may play a
significant role as bacterivores in some freshwater
plankton (Bird and Kalff, 1987; Sanders and Porter,
1988; Berninger et al., 1992). The importance of
mixotrophs as bacterivores varies considerably both
temporally and spatially, and the factors determining
their abundance and feeding rates in natural systems
are not entirely clear. However, light, temperature, dissolved nutrient concentrations, and food levels are
likely to affect both the abundance and the relative importance of phagotrophy versus photosynthesis in these
organisms (Bird and Kalff, 1987; Sanders et al., 1989,
1990; Jones and Rees, 1994). Many protozoa glean
bacteria from surfaces or attached to particles (Albright
et al., 1987; Caron, 1987), but the quantitative role of
bacterivorous protozoa in benthic habitats is relatively
unknown. Pioneering studies in this area include
Baldock and Sleigh (1988) and Bott and Kaplan
(1989). In experiments with stained lake sediment,
Starink et al. (1994) found over 90% of protozoa ingested bacteria at rates up to 73 bacteria protist1
hour1.
Small protozoa or “heterotrophic nanoplankton”
(flagellates, amoebas, and some small ciliates) capture
and consume bacteria individually, with rates of ingestion of tens to hundreds per hour (e.g., Fenchel,
1982b). Most ciliates, in contrast, use their complex
oral ciliature to collect hundreds into each food vacuole, producing feeding rates up to thousands of bacteria per hour (e.g., Taylor, 1986). Although the former
are likely to be selective in their feeding, the larger ciliates are probably very limited in their ability to discriminate. The use of fluorescent microspheres to measure bacterivory (McManus and Fuhrman, 1986)
resulted in renewed interest in the ability of protozoa
to discriminate against inorganic particles in favor of
bacteria (Nygaard et al., 1988) and in the influence of
chemical coatings and surface charge (Sanders, 1988).
Both ciliate and flagellate bacterivores discriminate
among bacteria on the basis of size (Sanders, 1988;
Simek and Chrzanowski, 1992; Simek et al., 1994).
Several studies have documented that ciliates differ in
their ability to use different bacteria as food. Some may
be unable to consume filamentous bacteria, while other
65
bacteria, especially pigmented ones, produce toxins
that render them unsuitable to support the growth of
some ciliates (Dive, 1973; Taylor and Berger, 1976).
A poorly understood and usually ignored facet of
bacterivory in protozoa is the role of extracellular products. The presence of some ciliates leads to the flocculation of bacteria (Hardin, 1943; Curds, 1963; Nilsson,
1976). Mucous-like extracellular materials are involved
in uptake of dissolved material and possibly small particles by pinocytosis in Amoeba. Small amoebas secrete
various hydrolases into their medium (Schuster, 1979)
as do flagellates. Porter (1988) suggested that flocculation and external digestion may be important in bacterivorous flagellates. The involvement of mucocysts or
other extrusomes is likely, but not confirmed.
As one might expect, feeding rate (functional response) and growth rate (numerical response) of
bacterivores increase with bacterial density, and the nature of these response curves has been studied because
of their relevance to interspecific competition and the
control of bacterial numbers. In addition to the expected hyperbolic functions, thresholds or sigmoid
responses have also been observed, and attributed to
changes from feeding to searching behavior at low particle concentrations (e.g., Taylor, 1978a; Sanders,
1988). Accordingly, low concentrations of bacteria
(104 – 107 per milliliter, usually 105 –106) can persist
even in stationary-phase cultures of bacterivorous protozoa. The behavioral and physiological mechanisms
responsible for this phenomenon are still in doubt
(Sambanis and Frederickson, 1988); however, it is
probably relevant to the densities of picoplankton observed in field situations.
7. Algivorous Protozoa
Larger protozoa are less likely to be bacterivores,
and most consume algae or other protozoa. Suspension-feeding ciliates each ingest a distinct size-spectrum
of particles, related to the form and function of the oral
apparatus (Fenchel, 1980), and this spectrum generally
shifts in larger ciliates toward larger particles, i.e.,
algae and other protozoa. However, even small prostomes can be important predators of flagellates
(Sommaruga and Psenner, 1993). Larger flagellates,
amoebas, and Heliozoa also consume other algae and
protozoa.
Protozoa feeding on relatively large items are more
likely to be selective, as the rate at which particles are
consumed is much reduced. For example, Rapport
et al. (1972) documented selective feeding by the heterotrich ciliate Stentor consuming other protozoa at up
to about five prey per minute. Stentors consistently
biased their feeding toward Tetrahymena relative to
FIGURE 13
(A) Peridinium; (B) Gymnodinium; (C) Gyrodinium; (D) Khawkinea halli; (E) Polytomella citri;
(F) Entosiphon sulcatum; (G) Petalomonas abcissa; (H) Peranema trichophorum; (I) Urceolus cyclostomas; (J)
Chilomonas paramecium; (K) Paraphysomonas vestita; (L) Spumella ( Monas) vivapara, two cell shapes; (M)
Ochromonas variabilisa; (N) Dinobryon sertularia(mixotrophic). (After: Bourrelly, 1968, L; Calaway and
Lackey, 1962, E, F, J, N; Eddy, 1930, A; Jahn and McKibben, 1937, D; Leedale, 1985, H; Pascher, 1913, M;
Lemmermann, 1914, K; Shawhan and Jahn, 1947, G; Smith, 1950, I.) Scale 5 m for E; 10 m for A, B, C,
G, I, J, K, L, M; 20 m for D, F, H, N.
66
3. Protozoa
three autotrophic flagellates when mixed versus single
species feeding suspensions were compared. The behavioral response of algivorous choreotrich ciliates appears
to involve both mechanical and chemical stimuli, and
suggests that although ciliates may ingest inert particles, they may discriminate among them as well as
among food organisms (Buskey and Stoecker, 1989).
Selective feeding has been noted for Amoeba (Mast and
Hahnert, 1935) and Thecamoeba (Page, 1977). Ciliates
may have relatively large ecological impacts on phytoplankton populations; in Lake Constance ciliates
consumed about 14% of the daily primary production
during a phytoplankton spring bloom, which was
equivalent to the amount removed by metazoan zooplankton (Weisse et al., 1990).
Although most phagotrophic flagellates are bacterivores, larger species may be effective algivores. Dinoflagellates, for example, may consume diatoms and
other large phytoplankton (see Section II.B.1).
Zoomastigina are probably under-rated as consumers
of algae (Canter, 1973; Goldman and Caron, 1985;
Suttle et al., 1986). The colorless chrysophyte Paraphysomonas (Fig. 2D) was observed to feed selectively
on diatoms, phagocytosing cells that were nearly their
own volume (Grover, 1989). The phagotrophic flagellate Peranema (Fig. 13H) feeds on related autotrophic
euglenids.
The consumption of filamentous algae (including
cyanobacteria) would appear to be mechanically difficult for protozoa, but many have specialized to this
role. Large amoebas, such as Pelomyxa include filamentous algae in their omnivorous diets, and some Testacealobosia, such as Arcella and Difflugia, are readily
cultured on filamentous algae which they engulf
through the aperture of their tests. The nassophorine
ciliates contain many forms which feed on filamentous
algae, including Nassula, Pseudomicrothorax, and
Frontonia. Nassula spp. (Fig. 11) appear to limit their
diet to Oscillatoria. Detailed ultrastructural studies
have revealed the role of the cyrtos in feeding, especially how it is used to break the algal filament so that
ingestion can begin (Tucker, 1968; Peck, 1985).
Abundant epibenthic diatoms, especially noncolonial and unattached pennate diatoms such as
Navicula, usually attract abundant herbivorous ciliates
and amoebas. Many dorsoventrally flattened and thigmotactic ciliates in the Phyllopharyngia and Nassophorea are diatom feeders. Their functional response
has been described by Balczon and Pratt (1996).
8. Histophagous Ciliates
Several genera and species of ciliates feed on metazoan tissue. Some, like Coleps hirtus and Prorodon
spp., are omnivores and only opportunistic histophages.
67
Strict histophages, feeding only on fresh tissue, include
all hymenostomes of the genus Ophryoglena (Fig. 10)
and some Tetrahymena spp. (e.g., Corliss, 1973; Lynn,
1975). Deltopylum is also a histophage; but in our
experience, it is relatively rare. Philaster appears to be
marine only. These forms feature complex life cycles
with reproductive cysts and multiple fission, which are
probably adaptations to an extremely patchy food resource (Section III.A). Like other carrion feeders, they
increase the efficiency of food webs by bypassing the
detrital pathway. Ophryoglena, and the related fish
parasite Ichthyophthirius, share the unique organelle of
Lieberkühn (Canella and Rocchi-Canella, 1976), or
watchglass organelle, whose function is unknown but
probably related to their feeding mode.
Related hymenostomes are insect parasites. Lambornella can be a parasite of the cuticle of mosquito
larvae (Washburn et al., 1988), and, it and some
Tetrahymena spp. may have significant impacts on
aquatic Diptera (Golini and Corliss, 1981; Egerter
et al., 1986). Espejoia feeds on the egg masses of
aquatic insects.
9. Metazoan Predators of Protozoa
Protozoa, as quantitatively important components
of plankton, aufwuchs, or sediment communities, probably contribute to the nutrition of many animal-feeding
guilds, such as filter-feeders, scrapers, and depositfeeders. However, because most protozoa are fragile
and without structures that resist digestion, they are seldom recorded among the gut contents of animals despite their importance (McCormick and Cairns, 1991).
Among benthic animals, minute predators, such as
Naididae (Oligochaeta), Orthocladinae (Diptera, Chironomidae), Turbellaria, and Copepoda consume protozoa selectively (Taylor, 1983b; Archbold and Berger,
1985; Kusuoka and Watanbe, 1989).
More is known about the feeding of planktonic
animals on protozoa (Porter et al., 1985; Sanders and
Wickham, 1993; Jack and Gilbert, 1997; Stoecker and
Capuzzo, 1990), reflecting the recent interest in the
microbial loop (Section III.C.3) and how picoplankton
production reaches higher trophic levels (e.g., Pomeroy,
1974; Sherr et al., 1987). Most planktonic rotifers consume heterotrophic flagellates, and many prey on ciliates (Arndt, 1993; Gilbert and Jack, 1993). Copepoda
especially are known to be consumers of protozoa, and
recent studies suggest that copepods may actively select
protozoa from among similar-sized plankton (Burns
and Gilbert, 1993; Wickham, 1995a,b). Cladocera can
ingest ciliates as well as both photosynthetic and
nonphotosynthetic flagellates (Sanders and Porter,
1988; Wickham and Gilbert, 1993). There is evidence,
however, that ciliates are not always suitable as food
68
W. D. Taylor and R. W. Sanders
for Cladocera (Debiase et al., 1990). Fish would seem
to be unlikely predators of protozoa, but protozoa are
of important food for some larval fishes (Kornienko,
1971; Kentouri and Divanach, 1986).
Many planktonic protozoa are capable of rapid or
saltatory movements that reduce their probability of
being captured. Speeds attained during these movements exceed 100 body lengths per second, and they
reduce the frequency of capture compared to species
that lack such movements (Gilbert, 1994). Morphological adaptations that reduce probablity of ingestion can
be induced by the presence of predators in only a few
hours in some ciliates (Wicklow, 1997). The ciliate
Lambornella reacts to the presence of predatory mosquito larvae by shifting morphologically and behaviorally to become a mosquito parasite (Washburn et al.,
1988).
energetics of protozoa (Section II) suggests that protozoa are more rather than less efficient in converting the
food they ingest into cytoplasm. Recent studies on
nutrient recycling by flagellates suggest that mineralization of nitrogen and phosphorus by protozoa may be
particularly low when it is most needed, that is, when
their food organisms are themselves N- or P-limited
(Goldman et al., 1985; Caron et al., 1988). In situ
measurements of nutrient release relative to ingestion
are few, but release of P for planktonic ciliates appears
to be close to 50%, or so we expect based on conversion efficiencies (Taylor and Lean, 1991). Protozoa
probably make an important contribution to the regeneration of nutrients in many environments, but no
more so than one would expect from their contribution
to consumption and production.
10. Protozoa and Nutrient Cycling
D. Evolutionary Relationships
Regeneration of the inorganic constituents of living
organisms occurs through a process involving ingestion, digestion and egestion by consumers, catabolism
and autolysis of all organisms, and the enzymemediated breakdown of nonliving organic matter by
decomposers. Autotrophs are net consumers of these
inorganic constituents, whereas phagotrophs regenerate
them. Decomposers, the heterotrophic bacteria and
fungi, regenerate carbon; but they may or may not be
net producers of other inorganic constituents, depending on the composition of the organic matter they
decompose. If it is poor in other constituents, such as
organic nitrogen and phosphorus (as when it is mostly
carbohydrate), then decomposers will compete with
autotrophs for these other constituents. In the final
analysis, the incorporation and regeneration of inorganic constituents will be in balance unless organic
matter is stored or exported.
Heterotrophic protozoa fit into this picture as typical consumers, eating other organisms and releasing
both unassimilated organic material and end products
of catabolism, and as major consumers of the decomposers. The latter role is manifested in the ability of
bacterivorous protozoa to accelerate the breakdown of
organic material by bacteria (Stout, 1980). This property of protozoa is mostly due to their keeping the bacteria in an actively growing state at moderate abundances, rather than directly due to their contribution to
the remineralization process.
Protozoa also contribute directly to mineralization,
and some early studies suggested protozoa were particularly active in releasing inorganic nutrients. However,
the basis of this view has been challenged (Taylor,
1986). Indeed, the relatively independent literature on
The relationship of protozoa to plants and animals, as well as the evolutionary relationships among
the protozoa, are controversial areas of current research. New tools for the study of phylogeny coming
from molecular biology are contradicting morphologybased classifications, but the number of organisms
examined is still small. Here, we will briefly introduce
some of the major theories and controversies.
One change in view which has occurred over
recent years is that the major dichotomy among living
things is not between plants and animals, but between
prokaryotes and eukaryotes. The evolution of eukaryotes from the prokaryotes is perhaps the major event in
organic evolution on earth, and may have occurred by
essentially Lamarckian means: the inheritance of
acquired characteristics (Taylor, 1987). That is, the
eukaryotic cell represents the symbiotic amalgamation
of several distantly related prokaryotes (Margulis,
1981). Although this concept is controversial, the controversy is largely one of degree in that the symbiotic
origin of some organelles, such as double-membraned
mitochondria and chloroplasts, is well supported by
molecular techniques and is now widely accepted.
Gupta et al. (1994) argued on the basis of a 70-kDa
heatshock protein that the eukaryotic nucleus (also
double-membraned) may also have been derived by
engulfing a bacterium.
Within the unicellular eukaryotes, or Protista, one
of the most primitive forms is the giant amoeba
Pelomyxa (Karyoblastea); it lacks mitochondria and
has amitotic nuclear fission. Many consider the flagellum to be as old as the eukaryote cell, but disagree
whether colorless flagellates are primitive or derived
from pigmented forms. Analyses of ribosomal RNA
3. Protozoa
strongly support the former argument for most
Zoomastigina. Some protists without mitochondria,
including the colorless (and parasitic) flagellate
Giardia, are clearly very close to the prokaryotes relative to other protozoa. This strongly suggests that their
features are primitive rather than derived (Sogin et al.,
1989, 1996). The same analysis reveals that the chlorophyte Chlamydomonas is more closely aligned with
higher plants than other protozoa, including other autotrophic phyla (Sogin et al., 1989).
Although our view of the relationship among unicellular eukaryotes is in a state of flux, the molecular
techniques currently being applied reveal that these
groups have had long and independent evolutionary
histories. For example, the genetic distances within
ciliates or flagellates are greater than the distances
between chlorophytes and higher plants, or between
plants and animals (Sogin and Elwood, 1986;
Gunderson et al., 1987).
There are two major schools of thought with respect to the relationship between protozoa and metazoa (Hanson, 1977). One considers that the protozoan
ancestor of metazoa is a colonial flagellate, probably a
choanoflagellate, and that Porifera, Placozoa, Cnidaria,
some gastrotrichs, and other animals with monociliated
epidermal cells are among the most primitive animals.
Others believe that primitive animals were derived by
the compartmentalization of a ciliated, multinucleate
protozoan. The early development of animals best supports the former hypothesis, as do the data collected
thus far on ribosomal RNA; the protozoa least distant
from animals are flagellates, not ciliates, which seem to
have had a very long independent history.
Perhaps the most important conclusions an ecologist can draw from the dynamic field of protist systematics is that although the protozoa are all unicellular or
colonial and comprise one to many phyla in different
classification schemes, they are more distant among
themselves than multicellular animals from the evolution point of view, and at least as morphologically and
physiologically diverse.
IV. COLLECTING, REARING, AND PREPARATION
FOR IDENTIFICATION
The small size, delicacy, and lack of hard parts in
most protozoa makes the quantitative collection and
enumeration of protozoa particularly difficult. The
methods for sample collection and examination vary
with the group of interest and the habitat, and there
is room for improvement in almost all the procedures
mentioned below. General techniques for the beginner
69
are provided in Lee et al. (1985) and Finlay et al.
(1988).
For some groups — especially amoebas and heterotrophic flagellates, when identification is required
or when the sample contains sediment or some other
material — examination of live organisms in fresh collections under a dissecting and/or compound microscope is the only method available. With care, this
can be done in a quantitative manner (e.g., Bott and
Kaplan, 1989). The “most probable number” method
of enumeration, based on inoculating culture medium
with small aliquots of sample and scoring the fraction of such inocula producing cultures, is common
in the study of soil protozoa and has been applied to
freshwater. However, the resultant estimates sometimes have been very low relative to direct counts
(Bott and Kaplan, 1989). Given the difficulty in culturing many freshwater protozoa, this method should
be reserved for estimating species known to be able
to grow on the medium used from small inocula. Extraction procedures have been devised for benthic
protozoa, but are not widely used (Hairston and
Kellerman, 1964; Fenchel, 1967). Fixed samples are
not useful in benthic studies, as the task of enumerating fixed but motionless protozoa in sediment is virtually impossible.
Studies on epibenthic or aufwuchs protozoa have
generally resorted to artificial substrates. Those that allow for microscopic examination are particularly useful: glass slides (Schönborn, 1977), petri dishes (Spoon
and Burbank, 1967; Taylor, 1983b), and strips of nylon
mesh (Taylor, 1983a). Polyurethane foam has been extensively used as a substrate in studying protozoan
community interactions and response to pollution
(Cairns et al., 1979).
Planktonic samples present fewer problems. Large
protozoa can be settled from samples and counted by
inverted microscopy as for phytoplankton. Useful fixatives are Lugol’s Iodine (1% v/v, or higher concentration in saline or hard water, e.g., Taylor and Heynen,
1987) or mercuric chloride (5% saturated mercuric
chloride plus a drop of 0.04% bromothymol blue, Pace
and Orcutt, 1981). These procedures do not lend themselves to the identification of ciliates, nor to the detection of chromatophores in small flagellates. Filtration
methods may aid with both of these problems. The
quantitative protargol or QPS method of Montagnes
and Lynn (1987a,b) and Skibbe (1994) produces permanent, quantitative, stained preparations for identification of ciliates and flagellates, although it requires
that samples be fixed in a concentrated Bouin’s fixative.
The use of epifluorescence microscopy on filtered samples (Caron, 1983) can facilitate the separation of
70
W. D. Taylor and R. W. Sanders
heterotrophic and phototrophic protists and is generally the method of choice for enumerating very small
planktonic protozoa, even though it does not lend itself
to identification.
Culture of most protozoa is best begun by maintaining samples collected in the field. Samples kept
in the laboratory will usually undergo succession, with
different species waxing and waning in abundance.
Screening out metazoan predators, such as copepods,
may encourage protozoa to increase in number.
Supplementing the organic or nutrient content of the
sample may help. Cereal grains (rice or wheat, boiled
for quicker effect or plain), broth from boiled vegetable
matter (hay, lettuce), or nutrient broth are commonly
used. Remember that adding too much is easy, and results in foul conditions that may encourage species different from those important in the original sample; or
one may simply kill all the aerobic protozoa. Generally,
additions of organic matter to cultures should not be
more than about 0.1% w/v for whole vegetable matter,
and no more than a tenth as much (100 mg/L) for more
labile material.
The first step in isolating species of interest is to
prepare some protozoa-free medium by filtering or
boiling water from the site where the protozoa were
collected, and inoculating individuals of the desired
species back in. Mineral water is frequently used for
culturing heterotrophic forms. Again, rice or wheat
grains or organic infusions are useful in promoting the
moderate growth of bacteria for bactivorous species.
Algae or other protozoa may be required as food for
others. White worms (enchytreids) are convenient food
for histophagous forms.
Ultimately, you may wish to establish monoxenic
(the organism of interest and its prey only) or axenic
cultures, possibly cloned from a single individual. Artificial freshwater media are described in Finlay et al.
(1988) and Thompson et al. (1988). Although involving marine protozoa, the recent efforts by Gifford
(1985) might prove a useful template for attempts to
culture difficult freshwater forms. A manual of elementary techniques is found in Finlay et al. (1988). General
and specific techniques for studying the ecology of freeliving protozoa are also compiled in Kemp et al.
(1993).
V. IDENTIFICATION OF PROTOZOA
A. Notes about the Key
It is often difficult and time consuming to identify
protozoa to the level of species. In many cases,
unambiguous identification requires specialized staining techniques or the use of electron microscopy. The
taxonomic grouping used in this key is an amalgamation of publications by specialists on the different
groups (e.g., Page, 1998; Lynn and Small, 1997; Lee
et al., 1985; Margulis et al., 1989). While the taxonomy is based on ultrastructural features and morphometric attributes that in many cases are not readily discerned with light microscopy, this key is designed to
make possible the identification of many protozoa to a
family or genus level using light microscopy alone.
Although observation of living organisms is important
for identification, the key should still be useful for
many fixed samples. Most of the specialized terms used
in the key are defined in the general glossary. If one
becomes more interested in identifying protozoa to a
species level, specialized techniques and keys in the
publications cited earlier can be used.
The genera listed in the key are representative of
the families and are not complete. Likewise, the illustrations used as examples here are of one or more
species considered typical of a genus, but do not necessarily represent all of the species in the genus.
Advances in the systematics of the protozoa are
continuous. Due to the heterogeneity of the group, it is
often difficult to remain abreast of the changes. A new
edition of Lee et al. (1985), An Illustrated Guide to the
Protozoa, is expected to be published by the Society of
Protozoologists at any time, and will likely have
changes in systematic organization.
Classification below the class level in the
Zoomastigina and the ordinal level in Heliozoa are not
conclusively agreed upon, or are under revision. Thus,
the use of familial names was avoided for these groups
in the key.
Phyla of flagellates marked with an asterisk (*)
contain many or mostly photosynthetic members;
genera listed have one or more species that lack chloroplasts and are thus heterotrophic, or contain chloroplasts, but also exhibit heterotrophic nutrition.
B. Taxonomic Key to Functional Groups of Protozoa
1a.
Locomotion via pseudopodia as in Figure 8. Rarely, flagella may be present as well (e.g., Fig. 9 C) ....................................................
Amoeboid Protozoa (Section V.D)
1b.
Simple cilia or compound ciliary organelles characteristic (but see Fig. 19A – P) and present in at least one part of life cycle; subpellicular infraciliature present even when cilia are not; two types of nuclei (macronucleus and micronucleus) with rare exceptions
(Fig. 23J). Ciliated Protozoa, Phylum Ciliophora (see Section V.E)
3. Protozoa
1c.
71
Commonly 1 to 4 flagella, 16 to 24 in one free-living genus (Paramastix); binary fission................................................... Flagellated
Protozoa (see Section V.C)
C. Taxonomic Key to Major Groups and Common
Genera of Flagellated Protozoa
1a.
With median groove (annulus and sulcus); two flagella, one extending transversely around the cell...............................................
Phylum Dinoflagellata*
(Peridinium, Gymnodinium, Gyrodinium; Fig. 13A – C)
1b.
Without a median groove .................................................................................................................................................................. 2
2a (1b).
Often large flagellates with pharyngeal rods (sometimes difficult to see) or containing the reserve material paramylon; one or usually
two flagella arise from within an anterior invagination (reservoir); contractile vacuole associated with reservoir; elongate; body
shape often plastic when living. Phylum Euglenida* (Khawkinea, Entosiphon, Petalomonas, Peranema, Urceolus; Fig. 13D,F – I)
2b.
Cells not as above ..............................................................................................................................................................................3
3a (2b).
Flagellates with two or four equal flagella. Phylum Chlorophyta* (Polytoma, Polytomella; Fig. 13E)
3b.
Cells with two unequal flagella...........................................................................................................................................................4
4a (3b).
Small cells with a deep, subapical gullet; two nearly equal length flagella arise from gullet................................... Phylum Cryptophyta* (Chilomonas; Fig. 7C, Fig. 13J)
4b.
Cells without deep gullet ....................................................................................................................................................................5
5a (4b).
Typically with two unequal flagella; long flagellum usually directed forward during swimming, with short flagellum directed backwards if emergent; compact Golgi body frequently visible anterior to nucleus of larger cells; chrysolaminarin vescicle(s) often fill
posterior of cell; contractile vacuole usually present in extreme anterior end of cell; many species capable of simultaneous photosynthesis and phagocytosis. Phylum Chrysophyta* (Dinobryon, Monas, Ochromonas, Paraphysomonas, Uroglena; Fig 7B, 13K – N,
14A)
5b.
Cell not as above. Phylum Zoomastigina ...........................................................................................................................................6
6a (5b).
Single anterior flagellum encircled laterally by a tentacular, funnel-shaped collar; solitary or colonial; with or without theca. Class
Choanoflagellida (Codosiga, Desmarella, Sphaeroeca, Diploeca, Monosiga, Salpingoeca; Fig. 4A, 14B – H)
6b.
Without collar, or with indistinct collar..............................................................................................................................................7
7a (6b).
Like Choanoflagellida above, but with second trailing flagella attached to the base of lorica; protoplasmic collar rudimentory.
Bicoecid flagellates, affinities uncertain (Bicoeca; Fig. 14I)
7b.
Without protoplasmic collar...............................................................................................................................................................8
8a (7b).
Flagellates with two unequal length flagella, one trailing; ingestion by pseudopodia. (Cercomonas; Fig. 14K)
8b.
Not as above ......................................................................................................................................................................................9
9a (8b).
With one or usually two flagella arising from a flagellar pocket (depression); characteristically elongate or bean shaped. Unique
organelle, the kinetoplast, usually associated with the flagella. Class Kinetoplastida (Bodo, Cephalothamnium, Pleuromonas,
Rhynchomonas; Fig. 14J,L,N – P)
9b
Cells with one or two nucleus-flagella complexes (karyomastigonts) each with 1 – 4 flagella; no Golgi apparatus; when two karyomastigonts, mirrored symmetry of nuclei and flagella. Diplomonads (Hexamita; Fig. 14M)
D. Taxonomic Key to the Major Groups and Common
Genera of Protozoa with Pseudopodia
1a.
Large, cylindrical or ovoid multinucleate amoeba; bacterial symbionts; usually containing mineral particles; nonmotile flagella-like
extensions; oxygen poor habitats Phylum Karyoblastea. (one genus, Pelomyxa; Fig. 8B, Fig. 16V)
1b.
Pseudopodia hyaline, usually blunt, eruptive (Fig. 8A,C,F) but filiform process may occur. May be flagellated stages........................2
1c.
Hyaline, filiform pseudopodia (filopodia) sometimes branching, not anastomosing (Fig. 8E); no known flagellate stages or spores.
Class Filosea.....................................................................................................................................................................................21
1d.
Threadlike and delicate pseudopodia with finely granular appearance, often branching and anastomosing to form complex reticulum (reticulopodia, Fig. 8G). Test often present. Phylum Granuloreticulosea ...................................................................................27
1f.
Long slender axopods (Fig. 8D) radiating from cell; with or without skeletal elements. Phylum Heliozoa .......................................32
FIGURE 14 (A) Uroglena americana(mixotrophic); (B) Desmarella moniliformis; (C,D) Sphaeroeca volvox,
individual cell and colony; (E) Codosiga botrys; (F) Diploeca placita; (G) Salpingoeca fusiformis; (H) Monosiga
ovata; (I) Bicoeca lacustris; (J) Bodo caudatus; (K) Cercomonas sp.; (L) Cephalothamnion cyclopum; (M)
Hexamita inflata; (N,O) Pleuromonas jaculans, attached and amoeboflagellate forms; (P) Rhynchomonas nasuta. (After: Bourrelly, 1968, L; Calaway and Lackey, 1962, N, O, P; Lackey, 1959, B, F; Lee, 1985, K;
Pascher, 1913, A; Pascher, 1914, C, D, E, G, H, I, J, M.) Scale 2.5 m for P; 5 m for F, G, H, I, K, L; 10
m for A, B, C, J, M, N, O; and 20 m for E.
72
3. Protozoa
73
2a (1b).
Small, usually cylindrical amoebae, many with a flagellated stage (Fig. 9C). Phylum Rhizopoda, Class Heterolobosea, Order
Schizopyrenida ...................................................................................................................................................................................3
2b
No flagellated stages. Class Lobosea ..................................................................................................................................................4
3a (2a).
Amoeboid form cylindrical, monopodial, eruptive; temporary flagellate stages common; nucleolus divides to form polar masses in
mitosis...................................Vahlkampfiidae (Valkampfia, Naegleria; Fig. 15A – B)
3b
Nucleolus disintegrates during mitosis, nuclear membrane does not; single known freshwater species; commonly flattened; no flagellate stage known...................................Gruberellidae (Stachyamoeba; Fig. 15C)
4a (1b).
Lacking external test. Subclass Gymnamoebia ...................................................................................................................................5
4b.
Incompletely enclosed in a test or other flexible cuticle of microscales. Subclass Testacealobosia .....................................................13
5a (4a).
Cylindrical or flattened; flattened forms with regular outline; no trailing uroidal filaments, with rare exceptions; not strikingly
eruptive. Order Euamoebida ..............................................................................................................................................................6
FIGURE 15 (A) Vahlkampfia avaria; (B) Naegleria; (C) Stachyamoeba lipophora; (D) Thecamoeba sphaeronucleolus; (E) Vanella miroides; (F) Amoeba proteus; (G) Mayorella bigomma; (H) Vexillifera telemathalassa;
(I) Hartmanella vermiformis; (J) Chaos illinoisense; (K) Saccamoeba lucens; (L) Trichamoeba cloaca; (M) Echinamoeba exudans; (N) Acanthamoeba; (O) Filamoeba nolandi (P) Hylodiscus rubicundus. (After: Bovee, 1985,
A, B, C, D, H, I, J, M, N, O; Kudo, 1966, F; Page, 1988, E, G, K, L, P.) Scale 10 m for A, B, C, E, I, M;
15 m for H, N, O, P; 30 m for D, G, K, L; 50 m for F; and 100 m for J.
74
W. D. Taylor and R. W. Sanders
5b.
Usually flattened; frequent changes in shape typical; sometimes eruptive; subpseudopodia usually present, often
furcate ..............................................................................................................................................................................................11
6a (5a).
Without subpseudopodia....................................................................................................................................................................7
6b.
With subpseudopodia.........................................................................................................................................................................9
7a. (6a).
Cell body flattened .............................................................................................................................................................................8
7b.
Cell body subcylindrical ...................................................................................................................................................................10
8a (7a).
Cell usually oblong; cresent-shaped hyaline margin at anterior end; pellicle-like layer, with dorsum often wrinkled and/or ridged;
usually uninucleate; no cytoplasmic crystals. Thecamoebidae (Thecamoeba; Fig. 15D)
8b.
Body usually fan-shaped, oval, or spoon-shaped, with hyaline margin occupying up to half of length. Vannellidae (Vannella, Fig. 15E)
9a (6b).
Subpseudopodia hyaline, blunt, digitiform, usually from anterior hyaline margin; uninucleate; nucleolar material in central body.
Paramoebidae (Mayorella, Fig. 15G)
9b.
Few slender, conical, or linear subpseudopodia, from anterior hyaline margin or cell surface; uninucleate. Vexilliferidae (Vexillifera,
Fig. 15H)
10a (7b).
Most species polypodial; length usually more than 75 m; uni- or multinucleate; numerous cytoplasmic crystals. Amoebidae
(Amoeba, Chaos, Trichamoeba; Fig. 15F,J,L)
10b.
Cell monopodial, pseudopods rare; uninucleate with central nucleolus; cytoplasmic crystals in some; cysts common. Hartmannellidae (Hartmannella, Saccamoeba; Fig. 15I,K)
11a (5b).
Cell regularly discoid, flattened ovoid, or fan-shaped; usually broader than wide; postcentral granular mass, usually surrounded,
sometimes completely, by hyaline border with short subpseudopodia. Hyalodiscidae (Hyalodiscus; Fig. 15P)
11b.
Not as above ....................................................................................................................................................................................12
12a (11b). Cell flattened, broad and irregular in outline, though sometimes elongate during locomotion; slender tapering subpseudopodia,
sometimes furcate, produced from broad, hyaline lobopodium; often with small lipid globules; uninucleate. Order Acanthopodida,
Acanthamoebidae (Acanthamoeba; Fig. 15N)
12b.
Several to many fine, sometimes furcate subpseudopodia, finer than in Acanthamoebidae. Echinamoebidae (Echinamoeba, Filamoeba; Fig. 15M,O)
13a (4b).
Test more or less rigid with distinct aperture. Order Arcellinida.......................................................................................................11
13b.
Discoid or sometimes globose amoeba incompletely closed in a flexible tectum, no well defined aperature. Order Himatismenida,
Cochliopodiidae (Cochliopodium, Fig. 16A)
14a (13a). Pseudopods conical, clear, sometimes anastomosing. Suborder Phryganellina, Phryganellidae (Phryganella; Fig. 16B – C)
14b.
Pseudopods digitate and finely granular ...........................................................................................................................................15
15a (14b). Test membranous or chitinoid, pliable or rigid; no plates or scales, but may have attached debris. Suborder
Arcellina...........................................................................................................................................................................................16
15b.
Test chitinoid or not, rigid, with embedded and/or attached plates, scales, siliceous granules. Suborder
Difflugina .........................................................................................................................................................................................18
16a (15a). Test flexible to semi-rigid; aperture ventral with variable shape. Microcoryciidae (Penardochlamys; Fig. 16H – I)
16b.
Test rigid, chitinoid ..........................................................................................................................................................................17
17a (16b). Test often areolar, smooth; aperture ventral, round. Arcellidae (Arcella, Pyxidicula; Fig. 16D,F – G)
17b.
Test oval to flask-shaped, nonareolar, clear; aperture terminal. Hyalospheniidae (Hyalosphenia; Fig. 16L – M)
18a (15b). Aperture round, broadly oval or wavy. Difflugiidae (Difflugia, Lesquereusia; Fig. 16J – K,N)
18b.
Aperture slit-like or narrowly oval ...................................................................................................................................................19
19a (18b). Aperture terminal .............................................................................................................................................................................20
19b.
Aperture antero-ventral, invaginated, slit-like with overhanging lip. Plagiopyxidae (Plagiopyxis; Fig. 16E)
20a (19a). Test particles rectangular. Paraquadrulidae (Quadrulella; Fig. 16O – P)
20b.
Test particles not rectangular. Nebelidae (Nebela; Fig. 16Q)
21a (1d).
Without distinct test, sometimes with scales. Order Aconchulinida ..................................................................................................22
21b.
With test. Order Testaceafilosida......................................................................................................................................................23
3. Protozoa
75
FIGURE 16
(A) Cochliopodium bilimbosum; (B, C) Phryganella nidulus; side and oral views; (D) Pyxidicula
operculata; (E) Plagiopyxis callida; (F, G) Arcella vulgaris, side and dorsal views; (H, I) Penardochlamys arcelloides, side and oral views; (J, K) Difflugia corona, side and oral views; (L, M) Hyalosphenia cuneata; (N) Lesquereusia spiralis; (O, P) Quadrulella symmetrica; (Q) Nebela collaris; (R) Penardia granulosa; (S) Chlamydophrys minor; (T, U) Lecythium hyalinum, dorsal and side; (V) Pelomyxa palustrus; (W) Pseudo-difflugia
gracilis; (After: Bovee, 1985, A, B, C, H, I, J, K, N, O, P, R, T, U; Deflandre, 1959, D, E, F, G, L, M, Q, S, W;
Kudo, 1966, V.) Scale 10 m for C, R, S; 25 m for A, H; 30 m for L, T, W; 45 m for N, O; 60 m for
Q; 90 m for B, E, F, J; and 500 m for V.
22a (21a). Filopodia nonanastomosing and more or less radiate. Vampyrellidae (Vampyrella; Fig. 17A)
22b.
Filopodia distally branching and anastomosing. Reticulosidae (Penardia; Fig. 16R)
23a (21b). Test without scales; may have spines, attached debris.......................................................................................................................24
23b.
Test with secreted, siliceous scales arranged in definitive patterns.....................................................................................................25
24a(23a).
Test round, thin; may have spines or spicules. Chlamydophryidae (Chlamydophrys, Lecythium; Fig. 16S – U)
24b.
Test not round. Pseudodifflugiidae (Pseudodifflugia; Fig. 16W)
76
W. D. Taylor and R. W. Sanders
FIGURE 17 (A) Vampyrella lateritia; (B) Paraeuglypha reticulata; (C) Euglypha tuberculata; (D, E) Trinema
enchelys, oral and side views; (F) Sphenoderia lenta; (G, H) Cyphoderia ampulla; (I, J) Campascus triqueter;
(K) Paulinella chromataphora; (L) Diplophyrys archeri; (M) Lieberkuehnia wagnerella; (N) Microcometes
paludosa; (O) Microgromia haeckeliana; (P) Biomyxa vagans; (Q) Chlamydomyxa montana; (R) Reticulomyxa
filosa. (After: Bovee, 1985, B, D, E, F, G, H, I, J, K, L, M, N, O, P, Q, R; Deflandre, 1959, A, C.) Scale 10
m for L, N, O; 15 m for K; 25 m for A, B, C; 40 m for D, F, G; 50 m for I, M, Q; 80 m for P; and
10,000 m for R.
25a (23b). Scales round to elliptical, thin, overlapping, adjacent or scattered. Euglyphidae (Paraeuglypha, Euglypha, Trinema, Sphenoderia;
Fig. 17B – F)
25b.
Scales not as above ...........................................................................................................................................................................26
26a (25b). Scales thick, cylindrical; test usually with aperture at end of a neck bent to one side. Cyphoderiidae (Cyphoderia, Campascus;
Fig. 17G – I)
26b.
Scales long, with long axes perpendicular to pseudostome; with two symbiotic cyanobacterial filaments. Paulinellidae (Paulinella;
Fig. 17K)
3. Protozoa
77
27a (1e).
Phylum Granuloreticulosea. With a test. Class Monothalamea ........................................................................................................28
27b.
Without a test. Class Athalamea...................................................................................................................................................... 30
28a (27a). Test with one pseudostome (opening)...............................................................................................................................................29
28b.
With more than one pseudostomes. Amphitremidae (Diplophyrys, Microcometes; Fig. 17L,N)
29a (28a). Test flattened on one side. Microgromiidae (Microgromia; Fig. 17O)
29b.
Test not flattened on one side. Lieberkuehnidae (Lieberkuehnia; Fig. 17M)
30a (27b). Naked amoebae with or without anastomosing pseudopodia; one or a few nuclei Order Athalamida..............................................31
30b.
Naked, multinucleate and highly reticulate plasmodia. Order Promycetozoida, Family Reticulomyxidae (Reticulomyxa; Fig. 17R)
31a (30a). Body mass changeable in form. Biomyxidae (Biomyxa; Fig. 17P)
31b.
Body mass more or less round and central. Chlamydomyxidae (Chlamydomyxa; Fig. 17Q)
32a (1f).
Class Heliozoa; Axopodia without visible axonemes........................................................................................................................33
32b.
Axonemes usually visible with light microscopy ...............................................................................................................................34
33a (32a). Cell enclosed in latticed organic capsule (skeleton); generally stalked; no centroplast; cell body spherical in adults. Order Desmothoracida (Clathrulina; Fig. 18A)
FIGURE 18
(A) Clathrulina elegans; (B) Actinophrys sol; (C) Actinosphaerium eichhorni; (D) Heterophrys
myriopoda; (E) Ciliophrys infusionum; (F) Acanthocystis turfacea; (G) Lithocolla globosa; (H) Raphidiophrys
elegans. (After: Deflandre, 1959, H; Kudo, 1966, B, C, D, E; Rainer, 1968, A, F, G.) Scale 15 m for E;
30 m for B, D, G; 50 m for A; 75 m for F, H; and 160 m for C.
78
W. D. Taylor and R. W. Sanders
33b.
Lack centroplast and apparently lack axonemes, the lack of microtubules (axonemes) in axopods has not been confirmed and this
order may be abandoned. Order Rotosphaerida (Lithocolla; Fig. 18G)
34a (32b). Stalked or not; long thin granule-studded axopods that usually arise from centroplast; axonemes frequently discernible with light
microscopy; usually with skeleton of siliceous or organic plates and/or spicules; highly contractile axopods or stalk. Order Centrohelida. (Heterophrys, Acanthocystis, Raphidiophrys; Figs. 7, 18D, F,H)
34b.
No skeleton ......................................................................................................................................................................................35
35a (34b). No centroplast or axoplast; axopods granual-studded and thicker at bases; large central nucleus surrounded by lacunar ectoplasm
or several nuclei at periphery of central area with vesicular ectoplasm. Order Actinophryida (Actinophrys, Actinosphaerium;
Fig. 18B – C)
35b.
Microtubule organizing center of dense plaques from the nuclear membrane or from centroplast; may be confused with
Actiniphryida without knowledge of fine structure (origin and pattern of axopod microtubules). Order Ciliophyrida (Ciliophrys;
Fig. 18E)
E. Taxonomic Key to Major Groups and Common Genera of Ciliates
1a.
Suctorial tentacles present; no true cytostome or cytopharynx; adults (trophonts) usually sessile, many species ectosymbiotic, some
planktonic species; cilia absent except in free-swimming dispersal stages; with or without lorica. Class Phyllopharygea, Subclass
Suctoria ..............................................................................................................................................................................................9
(Note: Patterns of division and release of the larvae form the basis for dividing the subclass into orders. Other characters are useful
in identification, but knowledge of the full life cycle is often required before suctorians can be confidently assigned to a genus.)
1b.
Cilia present in unencysted stages; no suctorial tentacles ....................................................................................................................2
2a (1b).
Conspicuous buccal ciliature at apical pole; buccal ciliature winds clockwise toward the center when viewed from oral end; somatic
cilature reduced or absent; mobile or sessile; solitary, gregarious or colonial; some species loricate; oral region can contract and
withdraw in most species. Class Oligohymenophora, Subclass Peritrichia ........................................................................................17
2b.
Not as above ......................................................................................................................................................................................3
3a (2b).
Oral area usually bordered by a well-developed adoral zone of membranelles (AZM) consisting of more than three membranelles
(polykineties); with or without ventral cirri ......................................................................................................................................26
3b.
Oral area not as above; no ventral cirri ..............................................................................................................................................4
4a (3b).
Cytostome at or near surface; buccal cavity, if present, without cilia or paroral membranes ..............................................................5
4b.
Cytostome at end of a buccal cavity with cilia or paroral membrane(s) associated .............................................................................6
5a (4a).
Cytostome at or near anterior end, or continuing down side as a slit ...............................................................................................52
5b.
Cytostome lateral or ventral ...............................................................................................................................................................7
6a (4b).
Paroral membrane(s) within or leading into buccal cavity ................................................................................................................65
(Sometimes difficult to see; follow both alternatives in key when in doubt)
6b.
No paroral membrane; simple cilia in buccal cavity .........................................................................................................................82
7a (5b).
Cytostome a barely visible lateral slit on the convex side of a tapering front end, or a lateral opening at the base of an anterior
proboscis..........................................................................................................................................................................................63
(see also Spathidiidae, 55)
7b.
Circular mouth located mid-ventrally; no proboscis; large cyrtos .......................................................................................................8
8a (7b).
Body nearlly ellipsoid, rounded in cross section, sometimes flattened ventrally; medium to large (some 100 m); densely ciliated
all over; oral depression present. Class Nassophorea, Order Nassulida, Nassulidae (Nassula; Fig. 20Z)
8b.
Body usually flattened; ventrum ciliated, dorsum bare or with a few cilia; anterior preoral arcs of right ventral ciliary rows continuous with more posterior parts. Class Phyllopharyngea, Order Cyrtophorida, Chilodonellidae (Chilodonella; Fig. 24T)
9a (1a).
Budding and cytokinesis on surface of trophont. Order Exogenida ..................................................................................................10
9b.
Budding begins in a pouch ...............................................................................................................................................................12
10a (9a).
Trophont basally attached to bottom of lorica near stalk. (Thecacineta; Fig. 19A)
10b.
Not as above ....................................................................................................................................................................................11
11a (10b). With vase-like lorica, sometimes with stalk-like base; tentacles extend through one or more slits. Urnulidae (Metacineta, Paracineta;
Fig. 19B – D)
3. Protozoa
79
FIGURE 19
(A) Thecacineta cothurniodes; (B, C) Metacineta mystacina, top and side views; (D) Paracineta
crenata; (E) Podophyra fixa, showing trophont, encysted form, and swarmer; (F) Acineta limnetis; (G) Sphaerophyra magna; (H) Trichophyra epistylidis; (I) Dendrocometes paradoxus; (J) Heliophyta reideri; (K) Tokophyra
quadripartita; (L) Multifasciculatum elegans; (M) Squalorophyra macrostyla; (N) Discophyra elongata; (O)
Stylocometes digitalis; (P) Dendrosoma radians. (After: Corliss, 1979, P; Goodrich and Jahn, 1943, F, K, L, M;
Kent, 1880 – 1882, G, I; Matthes, 1972, J, O; Noland, 1959, A, B, C, D, N; Small and Lynn, 1985, E, H.)
Scale 15 m for E, H, J, O; 30 m for A, D, F, G; 50 m for I, L, M, N; 75 m for B, K; and 200 m for P.
11b.
Trophont small, pyriform to spherical; usually stalked; tentacles apical or evenly distributed; often attached to other ciliates.
Podophryidae (Podophyra, Sphaerophyra; Fig. 19E,G)
12a (9b).
Budding and cytokinesis completed in brood pouch. Order Endogenida . . . ....................................................................................13
12b.
Cytokinesis begun in pouch, completed exogenously. Order Evaginogenida ....................................................................................15
13a (12a). Trophont without stalk or stalk-like process; attached to substrate by broad part of body. Dendrosomatidae (Dendrosoma,
Trichophyra; Fig. 19H,P)
80
W. D. Taylor and R. W. Sanders
FIGURE 20 (A) Hastatella radians; (B) Astylozoon faurei; (C) Urceolaria mitra; (D) Trichodina pediculis;
(E) Scyphidia physarum; (F) Cothurnia imberbis; (G) Vaginicola ingenita; (H, I) Zoothamnium arbuscula,
individual and colony; (J) Ophrydium eichhorni; (K) Vorticella campanula; (L) Pyxicola affinis; (M) Platycola
longicollis; (N) Thuricola folliculata; (O) Epistylis plicatilis; (P) Rhabdostyla pyriformis; (Q, R) Carchesium
polypinum, individual and colony; (S) Opercularia nutans; (T, U) Campanella umbellaria, individual and
colony; (V) Pseudomicrothorax agilis; (W) Microthorax pusillus; (X) Aspidisca costata; (Y) Euplotes patella;
(Z) Nassula ornata. (After: Corliss, 1959, V, Y; Kahl, 1930 – 1935, A, B, C, D, E, H, L, N, Q, R, T, U, W;
Kent, 1880 – 1882, I, J, K, O, S, X; Noland, 1959, F, G, M, P.) Scale 15 m for V, W; 20 m for A, B, G, P;
25 m for D, H, F, X; 30 m for C, Z; 40 m for E, L, M, N, S; 50 m for O, Y; 75 m for K, Q, U; and
200 m for J.
3. Protozoa
13b.
81
Not as above ....................................................................................................................................................................................14
14a (13b). With lorica, usually stalked; tentacles in a few fascicles or distributed over body. Acinetidae (Acineta; Fig. 19F)
14b.
Lacks lorica; tentacles single, fascicles, or evenly distributed. Tokophryidae (Tokophyra, Multifasciculatum; Fig. 12 and 19K,L)
15a (12b). Conical tentacles; body often hemispherical . Dendrocometidae (Dendrocometes, Stylocometes; Fig. 19I,O)
15b.
Not as above, capitate tentacles .......................................................................................................................................................16
16a (15b). Body discoidal, flattened against substrate, with peripheral attachment ring. Heliophryidae (Heliophrya, Fig. 19J)
16b.
Thin tentacles; body spherical to ovoid; some species loricate and/or stalked. Discophryidae (Discophrya, Squalorophyra;
Fig. 19M,N)
17a (2a).
Trophont mobile; symbiotic on other organisms; with complex, aboral holdfast. Order Mobilida...................................................18
17b.
Trophont only rarely motile, attached with stalk; often attached to other organisms; many gregarious or colonial species. Order
Sessilida............................................................................................................................................................................................19
18a (17a). Denticles of holdfast with hooks and spines; ectosymbionts on Hydra, fishes, Amphibia. Trichodinidae (Trichodina; Fig. 20D)
18b.
Denticles of holdfast simple, toothed; elongated macronucleus, ectosymbionts on turbellarians. Urceolariidae (Urceolaria; Fig. 20C)
19a (17b). Free-swimming; aboral end drawn to point with one or two rigid bristles or cilia; swim with oral end forward. Astylozoidae
(Astylozoon, Hastatella; Fig. 20A,B)
19b.
Aborally attached to substratum directly or by stalk ........................................................................................................................20
20a (19b). Colonial, with zooids in gelatinous matrix. Ophrydiidae (Ophrydium; Fig. 20J)
20b.
Not in gelatinous matrix ..................................................................................................................................................................21
21a (20b). Stalk absent, tapering body may resemble short stalk. Scyphidiidae (Scyphidia; Fig. 20E)
21b.
Stalked, or, if stalkless, in lorica .......................................................................................................................................................22
22a (21b). With contractile stalk; solitary or colonial........................................................................................................................................23
22b.
If stalked, stalk not contractile; solitary or colonial ..........................................................................................................................24
23a (22a). Each zooid independently contractile; if solitary, entire stalk contractile. Vorticellidae (Carchesium, Vorticella; Fig. 20K,Q – R)
23b.
Entire colony contracts in unison due to shared, continuous myonemes. Zoothamniidae (Zoothamnium; Fig. 20H – I)
24a (22b). With vase-shaped lorica, stalkless or short stalked; slender body attached to lorica aborally. Vaginicolidae (Cothurnia, Platycola,
Pyxicola, Thuricola, Vaginicola; Fig. 20F,G,L – N)
24b.
Not as above ....................................................................................................................................................................................25
25a (24b). Stalked; peristomal area raised on short neck with furrow separating it from margin of zooid. Operculariidae (Opercularia;
Fig. 20S)
25b.
Stalked; solitary or colonial; peristomal area not on neck, no furrow. Epistylididae (Campanella, Epistylis, Rhabdostyla; Fig. 20O,
P, T, U)
26a (3a).
Locomotor cirri present on ventral surface; generally dorso-ventrally flattened. Class Spirotrichea, subclass Hypotrichia................27
26b.
Not as above ....................................................................................................................................................................................36
27a (26a). Adoral zone of membranelles well developed, prominent .................................................................................................................28
27b.
AZM reduced; no marginal or caudal cirri, ventral cirri conspicuous. Class Spirotrichea, Subclass Hypotrichia, Order Euplotida,
Aspidiscidae (Aspidisca; Fig. 20X)
28a (27a). Right marginal row of cirri absent, one or more left marginal cirri; large caudal cirri; paroral membrane only as a field on right of
oral area. Class Spirotrichea, Subclass Hypotrichia, Order Euplotida (Euplotes; Fig. 20Y)
28b.
Left oral cilia as a “collar” and “lapel”, right oral cilia variable with one or more paroral membranes. Class Spirotrichea, Subclass
Stichotrichia .....................................................................................................................................................................................29
29a (28b). Ventral cirri not in distinct file from anterior to posterior (exception: Gastrostyla) Oxytrichidae (Gastrostyla, Oxytricha, Stylonychia; Fig. 21A,C,E)
29b.
Ventral cirri in distinct linear or zig-zag file(s) ..................................................................................................................................30
30a (29b). Frontoventral cirri run length of ventrum as zig-zag files. Urostylidae (Urostyla, Uroleptus; Fig. 21B,D)
82
W. D. Taylor and R. W. Sanders
30b.
Ventral cirri in one or more linear files of varied length....................................................................................................................31
31a (30b). Body in lorica; without marginal cirri; anterior end long, narrow, contractile, Chaetospiridae (Chaetospira; Fig. 21N,O)
31b.
Not as above ....................................................................................................................................................................................32
32a (31b). Files of cirri curved or spiralled along body......................................................................................................................................33
32b.
Files of cirri straight or oblique, ventral only....................................................................................................................................35
33a (32a). Files of cirri end on dorsum; oral region extends over one-fourth of body length. Strongylidiidae (Strongylidium; Fig. 21P)
33b.
Not as above ....................................................................................................................................................................................34
FIGURE 21 (A) Gastrostyla steini; (B) Uroleptus piscis; (C) Oxytricha fallax; (D) Urostyla grandis; (E) Stylonychia mytilus; (F) Gonostomum affine; (G) Amphisiella oblonga; (H) Stichotricha aculeata; (I) Hypotrichidium conicum; (J) Discomorphella pectinata; (K) Metopus es; (L) Myelostoma flagellatum; (M) Saprodinium
dentatum; (N, O) Chaetospira mülleri, contracted and extended forms; (P) Strongylidium crassum; (Q)
Psilotricha acuminata; (R) Caenomorpha medusula; (S) Tintinnidium fluviatile; (T) Tintinnopsis cylindricum;
(U) Strombidinopsis setigera; (V) Strombidium viride; (W) Halteria grandinella; (X) Strombilidium gyrans. (After: Jankowski, 1964a, b, J, M; Kahl, 1930 – 1935, F, G, H, I, K, L, N, O, P, Q, R, V, W, X; Kent, 1880 – 1882,
A, B, C, D, E; Noland, 1959, S, T, U.) Scale 15 m for L; 25 m for H, W, X; 30 m for F, I, J, P,Q, R, T;
40 m for G, K, M, N, O, S, U, V; 60 m for B, E; 80 m for A, C; and 140 m for D.
3. Protozoa
83
34a (33b). Ventral files of cirri curve to left rear. Psilotrichidae (Psilotricha; Fig. 21Q)
34b.
Ventral files of cirri spiral obliquely around body. Spirofilidae (Hypotrichidium, Stichotricha; Fig. 21H,I)
35a (32b). Files of cirri parallel to long axis of oral region along its right border. Gonostomatidae (Gonostomum; Fig. 21F)
35b.
Files of cirri not parallel to oral region, one or more of the ventral cirral files extends from anterior to well past mid-ventrum.
Amphisiellidae (Amphisiella; Fig. 21G)
36a (26b). Body surface ciliated ........................................................................................................................................................................46
36b.
Somatic ciliature sparse or absent.....................................................................................................................................................37
37a (36b). Body flattened, rigid, often with spines; body ciliature present as short, generally obvious rows; oral ciliature relatively inconspicuous; mainly anaerobic. Class Spirotrichea, Order Odontostomatida ................................................................................................38
37b.
Not as above ....................................................................................................................................................................................40
38a (37a). Body box-shaped, generally with short posterior spines; short rows of body cilia at front and rear, those in front parallel to and forward of oral opening. Epalxellidae (Saprodinium; Fig. 21M)
38b.
Body round to discoidal ...................................................................................................................................................................39
39a (38b). Distinct anterior spine; band of cilia on ridge overhanging buccal cavity. Discomorphellidae (Discomorphella; Fig. 21J)
39b.
Body ciliature very sparse; lacks spines; pair of cirri at posterior end. Myelostomatidae (Myelostoma; Fig. 21L)
40a (37b). Membranelles numerous in complete or almost complete circle at oral end; body generally conical or bell-shaped; mostly planktonic. Class Spirotrichea, Subclasses Choreotrichia and Oligotrichia................................................................................................41
40b.
Body rounded or conical, spirally twisted to left; twisting of body less prominent than Metopidae; often with long caudal spine; oral
region spiralled; dense preoral cilia along anterior border of oral cavity; body cilia absent except for caudal tuft and anterior cirruslike tufts; anaerobic. Class Spirotrichea, Order Armophorida (Metopus, Caenomorpha; Fig. 21M,R)
41a (40a). Oral ciliature forms a closed circle. Subclass Choreotrichia..............................................................................................................42
41b.
Oral cilature forms an open anterior collar with an anterioventral “lapel”. Subclass Oligotrichia ...................................................45
42a (41a). Loricate, attached to inner wall of lorica ......................................................................................................................................... 43
42b.
Not loricate ......................................................................................................................................................................................44
43a (42a). Delicate, gelatinous or mucoid tubular lorica, with attached debris, often translucent . Tintinnididae (Tintinnidium; Fig. 21S)
43b.
Lorica rigid, agglomerate of mineral grains and diatom fragments. Codonellidae (Tintinnopsis; Fig. 21T)
44a (42b). Rows of somatic ciliature equally distributed around body, extending length of body; body cilia may be long. Strombidinopsidae
(Strombidinopsis; Fig. 21U)
44b.
One or more usually short rows of body cilia; body ciliature short. Suborder Strobilidiina, Strobilidiidae (Strobilidium, Fig. 21X)
45a (41b). Somatic ciliature reduced to a few bristle-like cirri arranged circumferentially around the equator effecting a jumping motion. Halteriidae (Halteria; Fig. 21W)
45b.
Somatic ciliature greatly reduced or absent, no bristles; sometimes with prominent girdle close to middle of cell. Strombidiidae
(Strombidium; Fig. 21V)
46a (36a). Uniform somatic ciliation; often with conspicuous tuft of caudal cilia; buccal membranelles large, but not always obvious; anterior
end of body twisted left; oral region spiralled with dense preoral ciliation; in richly organic environments with low oxygen. Class
Spirotrichea, Order Armophorida, Metopidae (Metopus; Fig. 21K)
46b.
Not as above ....................................................................................................................................................................................47
47a (46b). Somatic ciliature well developed, uniform, usually inserts on oral region; body size large; many contractile species; rarely loricate;
some species pigmented. Class Heterotrichea, Order Heterotrichida ................................................................................................48
47b.
Somatic ciliature well developed; body large (500 –1,000 m), broad; buccal cavity prominent, funnel-like; macronucleus elongate;
carnivorous. Class Colpodea, Order Bursariomorphida, Bursariidae (Bursaria; Fig. 22C)
48a (47a). Body trumpet-shaped; highly contractile; oral ciliature spirals clockwise around flared anterior end; often pigmented and/or with algal endosymbionts. Stentoridae (Stentor; Fig. 22F)
48b.
Body not trumpet-shaped .................................................................................................................................................................49
49a (48b). Body laterally compressed, pyriform to elipsoid; often pigmented pink, red, or purple; long, narrow peristome on left margin;
noncontractile. Blepharismidae (Blepharisma; Fig. 22B)
FIGURE 22
(A) Spirostomum minus; (B) Blepharisma lateritium; (C) Bursaria truncatella; (D) Climacostomum virens; (E) Condylostoma tardum; (F) Stentor polymorphus, half extended; (G) Actinobolina radians; (H)
Coleps hirtus; (I) Bryophyllum lieberkühni; (J) Metacystis recurva; (K) Lacrymaria olor; (L) Askensia volvox;
(M) Urotricha farcta; (N) Mesodinium pulex; (O) Vasicola ciliata; (P) Trachelophyllum apiculatum; (Q)
Enchelyodon elegans; (R) Homalozoon vermiculare; (S) Enchelys simplex; (T) Chaena teres; (U) Spathidium
spathula; (V, W) Didinium nasutum, live and silver stained. (After: Dragesco, 1966a, K, S, V, W; Dragesco,
1966b, P, R; Kahl, 1930 – 1935, A, B, D, E, F, G, H, I, J, L, M, N, O, Q, T, U; Kent, 1880 – 1882, C.) Scale
10 m for M, N; 20 m for H, J, L, P, S; 30 m for G, O, U; 40 m for B, K, T; 60 m for E, Q, R; 80 m for
D, V, W; 100 m for A, F, I; and 200 m for C.
84
3. Protozoa
49b.
85
Body shape not as above ..................................................................................................................................................................50
50a (49b). Body large, very long; oral cavity shallow; peristomal area long, narrow, oral ciliature sometimes inconspicuous; contractile. Spirostomidae (Spirostomum; Fig. 22A)
50b.
Buccal ciliature prominent; oral cavity deeper than above ................................................................................................................51
51a (50b). Buccal cavity occupying much of anterior part of body; paroral membrane inconspicuous; somatic ciliation dense; with or without
endosymbiotic algae. Climacostomidae (Climacostomum; Fig. 22D)
51b.
Paroral membrane prominent; contractile; somatic ciliation dense. Condylostomatidae (Condylostoma; Fig. 22E)
52a (5a).
Body ovoid; with tentacles extending from the cell in all directions when at rest, but retracted and hardly visible when swimming.
Class Litostomatea, Order Pharyngophorida, Actinobolinidae (Actinobolina; Fig. 22G)
52b.
Tentacles absent ...............................................................................................................................................................................53
53a (52b). Translucent CaCO3 plates (armor) in cortex; typically barrel-shaped; body frequently spiny, often with prominent anterior and caudal thorns; long caudal cilium common; brosse present, but inconspicuous. Class Prostomatea, Order Prorodontida, Colepidae
(Coleps; Fig. 22H)
53b.
No such armor, not as above ............................................................................................................................................................54
54a (53b). Sessile in pseudochitinous lorica; one or more caudal cilia; paratenes obvious. Class Prostomatea, Order Prostomatida, Metacystidae (Metacystis, Vasicola; Fig. 22J,O)
54b.
Not as above ....................................................................................................................................................................................55
55a (54b). Mouth at anterior end, rounded or only slightly elongated ..............................................................................................................56
55b.
Apex fan-shaped to varying degree; mouth a long slit, beginning at anterior end; slit may extend down side of cell. Litostomatea,
Order Haptorida, Spathidiidae (Byrophyllum, Spathidium; Fig. 22I,U)
56a (55a). Mouth at anterior pole, surrounded by an unciliated area and often raised as a blunt cone; circumferential ciliary girdle of closely
apposed cilia, otherwise naked or with short cilia. Class Litostomatea, Order Haptorida ................................................................57
56b.
Uniform ciliation around mouth, and typically over the entire cell ...................................................................................................58
57a (56a). Girdle ciliature of one type, one or two girdles. Didiniidae (Didinium; Figs. 6, 22V,W)
57b.
Body ciliature of two types, one cirrus-like, as a single girdle. Mesodiniidae (Askenasia, Mesodinium; Figs. 22L,N)
58a (56b). Rear one-third to one-fifth of body unciliated, except for one or more long caudal cilia; other somatic ciliation evenly distributed.
Class Prostomatea, Order Prorodontida, Urotrichidae (Urotricha; Fig. 22M) ......................................................................................
58b.
Not as above ....................................................................................................................................................................................59
59a (58b). Body uniformly ciliated at rear; mouth with oral dome or bulb; cytopharynx not permanently inverted, but inverts during ingestion.
Class Litostomatea, Order Haptorida ..............................................................................................................................................60
59b.
Not as above; lack oral dome ...........................................................................................................................................................62
60a (59a). Anterior tapering to a ciliated neck; neck set off by groove and a circle of longer cilia; contractile . Lacrymariidae (Lacrymaria;
Fig. 22K)
60b.
Not as above ....................................................................................................................................................................................61
61a (60b). Body long, generally 4 times width; ovoid or flask-shaped; oral region simple dome, sometimes pointed. Trachelophyllidae
(Chaenea, Trachelophyllum; Fig. 22P,T)
61b.
Oral region flattened at apex; cytostome circular to ovoid; body usually shorter than four times width. Enchelyidae (Homalozoon,
Enchelys, Enchelyodon; Fig. 22Q,R,S)
62a (59b). Short “dorsal brush” (brosse) extends backward from apical mouth on one side (may not be obvious without silver staining); body
ellipsoid. Class Prostomatea, Order Prorodontida, Prorodontidae (Prorodon, Pseudoprorodon; Fig. 23A,B)
62b.
Dorsal brush lacking, may have reduced brosse; similar to, and easily confused with Prorodon. Class Prostomatea, Order Prorodontida, Holophryidae (Holophyra; Fig. 23C)
63a (7a).
Body long, flattened; mouth on convex side of tapering anterior end; ciliated only on right side of body. Class Karyorelictea, Order
Loxodida, Loxodidae (Loxodes; Fig. 23J)
63b.
Not as above ....................................................................................................................................................................................64
64a (63b). Cytopharynx permanent, round, located at base of proboscis or tapering front end; ciliature along proboscis; toxicysts on ventrum
of proboscis. Class Litostomatea, Order Pharyngophorida, Tracheliidae (Dileptus, Paradileptus, Trachelius; Fig. 23D,E,H)
FIGURE 23
(A) Prorodon teres; (B) Pseudoprorodon ellipticus; (C) Holophyra simplex; (D) Trachelius
ovum; (E) Paradileptus robustus; (F) Amphileptus claparedi; (G) Litonotus fasciola; (H) Dileptus anser; (I)
Loxophyllum helus; (J) Loxodes magnus; (K) Cyrtolophosis mucicola; (L, M, N) Philasterides armata, live,
silver stained, and oral detail of silver-stained specimen; (O) Loxocephalus plagius; (P) Urozona bütschlii;
(Q) Balanonema biceps; (R) Pleuronema coronatum; (S) Histiobalantium natans; (T) Cohnilembus pusillus;
(U) Uronema griseolum; (V) Cinetochilum margaritaceum; (W) Cyclidium glaucoma; (X) Calyptotricha
pleuronemodies. (After: Corliss, 1959, R; Dragesco, 1966a, I; Grolière, 1980, M, N; Kahl, 1930 – 1935, A, B,
C, F, G, J, K, O, P, Q, S, V, W, X; Kudo, 1966, D, E, H; Noland, 1959, L, T, U.) Scale 10 m for K, Q; 15
m for P, V; 20 m for T, U, W, X; 25 m for G, H, L, M; 30 m for C, I, S; 40 m for B, R; 50 m for F; 60
m for A, O; and 75 m for D, E, J.
86
3. Protozoa
87
64b.
Mouth slit-like, located along convex side of tapering anterior end, evident only when feeding; body laterally compressed; both
sides ciliated, cilia may be only bristles on 1 side. Class Litostomatea, Order Pleurostomatida, Amphileptidae (Amphileptus, Litonotus, Loxophyllum; Fig. 23F,G,I)
65a (6a).
With linear oral furrow or groove leading from anterior end to mouth, bordered on right by paroral membrane ............................66
65b.
Without linear preoral groove bordered by membranes, but generally with membranes inside oral cavity .......................................71
66a (65a). With an apparent double paroral membrane on the right side of the furrow (actually the paroral membrane plus a row of somatic
cilia). Class Oligohymenophora, Order Scuticociliatida, Cohnilembidae (Cohnilembus; Fig. 23T)
(Many genera of the Order Scuticociliatida, e.g., lines 66 – 70 and lines 75 – 76, are difficult to identify without special preparation
techniques such as Protargol or other silver staining)
66b.
Paroral membrane along furrow not “double”.................................................................................................................................67
67a (66b). Preoral furrow long, shallow, ciliated with three ciliary fields; generally a single caudal cilium; body long, ovoid. Class Oligohymenophora, Order Scuticociliatida, Philasteridae (Philasterides; Fig. 23L – N)
67b.
Bottom of preoral furrow not ciliated, paroral membrane curves around rear of mouth ..................................................................68
68a (67b). Dwells in lorica that is open at both ends; resembles Pleuronema or Cylidium. Class Oligohymenophora, Order Scuticociliatida,
Calyptotrichidae (Calyptotricha; Fig. 23X)
68b.
Not in a lorica as above ...................................................................................................................................................................69
69a (68b). Small (15 – 60 m long); with few somatic cilia; distinct caudal cilium. Oligohymenophora, Order Scuticociliatida, Cyclidiidae
(Cyclidium; Fig. 23W)
69b.
Larger than above ............................................................................................................................................................................70
70a (69b). Paroral membrane a prominent velum, extends from anterior to well past equator of cell; one to many stiff, long caudal cilia; a single contractile vacuole. Class Oligohymenophora, Order Scuticociliatida, Pleuronematidae (Pleuronema; Fig. 23R)
70b.
Somatic ciliation uniform; long stiff cilia distributed over body surface; paroral membrane less prominent than above. Class Oligohymenophora, Order Scuticociliatida, Histiobalantiidae (Histiobalantium; Fig. 23S)
71a (65b). Oral cavity huge, covers most of ventrum; dorsal side convex; cilia of paroral membrane long. Class Nassophorea, Order Peniculida, Lembadionidae (Lembadion; Fig. 24G)
71b.
Mouth smaller than above, not over half of body length ..................................................................................................................72
72a (71b). With one or more long caudal cilia...................................................................................................................................................73
72b.
Without caudal cilia .........................................................................................................................................................................77
73a (72a). With one or two girdles of cilia ........................................................................................................................................................74
73b.
Cilia not in girdles, usually in longitudinal rows ..............................................................................................................................75
74a (73a). Oral cavity equatorial; distinct constriction in the ciliated girdle; ends bare except for caudal cilium. Class Oligohymenophora, Order Scuticociliatida, Urozonidae (Urozona; Fig. 23P)
74b.
Oral cavity deep, equatorial; two distinct ciliary girdles; ciliary tuft on posterior end. Class Nassophorea, Order Peniculida, Urocentridae (Urocentrum; Fig. 24D)
75a (73b). Mouth at, or anterior to, middle of cell ............................................................................................................................................76
75b.
Oral area large, toward posterior half of cell (⬇ mid-ventral); body usually flattened; cilia denser ventrally. Class Oligohymenophora, Order Scuticociliatida, Cinetochilidae (Cinetochilum; Fig. 23V)
76a (75a). Small, 50 m; anterior pole flat, unciliated. Class Oligohymenophora, Order Scuticociliatida, Uronematidae (Uronema; Fig. 23U)
76b.
Somatic ciliation even; body long-ovoid; oral area small, closer to anterior end. Class Oligohymenophora, Order Scuticociliatida,
Loxocephalidae (Balanonema, Loxocephalus; Fig. 23O,Q)
77a (72b). Body small, ovoid to ellipsoid; in mucilaginous case from which it can emerge freely; cytostome near anterior end. Class Colpodea,
Order Cyrtolophosidida, Cyrtolophosidae (Cyrtolophosis; Fig. 23K)
77b.
Not associated with mucilaginous envelope .....................................................................................................................................78
78a (77b). Body distinctly heart- or cone-shaped; oral region relatively large, covers most of ventral surface. Class Nassophorea, Order Peniculida, Stokesiidae (Stokesia; Fig. 24B)
78b.
Body shaped otherwise .....................................................................................................................................................................79
79a (78b). Body ovoid to ellipsoid; mouth in anterior half of body, pointed in front, with pre- and/or post-oral sutures, sometimes hard to see;
88
W. D. Taylor and R. W. Sanders
FIGURE 24 (A) Frontonia leucas; (B) Stokesia vernalis; (C) Glaucoma scintillans; (D) Urocentrum turbo;
(E) Disematostoma bütschlii; (F) Turaniella vitrea; (G) Lemabadion magnum; (H) Colpidium colpoda;
(I) Paramecium caudatum; (J) Clathrostoma viminale; (K, L) Maryana socialis, individual and colony; (M) Plagiopyla nasuta; (N) Bresslaua vorax; (O) Tetrahymena pyriformis; (P, Q) Tillina magna, live and line drawing
of silver-stained specimen; (R, S) Colpoda steinii, live and silver stained; (T) Chilodonella uncinata. (T) (After:
Corliss, 1959, O, R; Dragesco, 1966b, B; Kahl, 1930 – 1935, A, C, D, E, F, G, H, I, J, K, L, M, P; Kudo, 1966,
N; Lynn, 1976, S; Lynn, 1977, Q; Noland, 1959, T.) Scale 15 m for G. O. R; 25 m for C, H; 30 m for
D, F; 40 m for B, E, J, M; 60 m for I, N; and 75 m for A, K, Q.
3. Protozoa
89
nematodesmata prominent to side and rear of mouth; contractile vacuoles with long collecting canals. Class Nassophorea, Order
Peniculida, Frontoniidae (Disematostoma, Frontonia; Fig. 24A)
79b.
Mouth typically rounded or blunt in front; body ciliature with preoral suture, no postoral suture; one paroral membrane and three
membranelles, which are typically inconspicuous. Order Hymenostomatida, Suborder Tetrahymenina ...........................................80
(Genera of this suborder, lines 80 and 81, are difficult to identify without special preparation techniques, i.e., silver staining)
80a (79b). Body pyriform to cylindrical; bases of membranelles of uniform width; mouth roughly triangular. Tetrahymenidae (Tetrahymena;
Fig. 1, 24O)
80b.
Not as above ....................................................................................................................................................................................81
81a (80b). Right ventral cilia curve left and twist forward parallel to suture. Turaniellidae (Colpidium, Turaniella; Fig. 24F,H)
81b.
Right ventral cilia curve left, but do not twist forward toward mouth. Glaucomidae (Glaucoma; Fig. 24C)
82a (6b).
In a gelatinous tube-like lorica, which may have dichotomous branches; with a terminal cone-like protuberance bearing long cilia
and surrounded by a circular adoral groove leading to the buccal cavity. Class Colpodea, Order Colpodida, Marynidae (Maryna;
Fig. 24K,L)
82b.
No gelatinous case; mouth area not as described above ...................................................................................................................83
83a (82b). Body small (100 m), laterally flattened; ciliation relatively sparse, on pellicular ridges................................................................84
83b.
Not as above ....................................................................................................................................................................................85
84a (83a). Cytostome in rear half of body; small cyrtos hidden by ventral pellicular fold. Class Nassophoria, Order Microthoracida, Microthoracidae (Microthorax; Fig. 20W)
84b.
Cytostome opens laterally in anterior one-third of body, long tubular cyrtos; body nearly oval. Class Nassophoria, Order Propeniculida, Leptophrygidae (Pseudomicrothorax; Fig. 20V)
85a (83b). Body ciliation uniform; ventral cytostome in an oral vestibule; nematodesmata form a ring or “basket” around mouth; similar to
Frontoniidae. Class Nassophorea, Order Peniculida, Clathrostomatidae (Clathrostoma; Fig. 24J)
85b.
Mouth area sunken in, with oral groove leading to it, or with ciliated pharyngeal tube, or both......................................................86
86a (85b). Medium to large (150 m in some species); body foot-shaped or ellipsoidal, with pointed or rounded caudal end; uniform body
ciliation; long, broad, ciliated oral groove leads into buccal cavity; two contractile vacuoles; one species with algal symbiont. Class
Nassophorea, Order Peniculida, Parameciidae (Paramecium; Fig. 6 and 24I)
86b.
Not as above; with single contractile vacuole ...................................................................................................................................87
87a (86b). Oral groove transverse, in anterior part of body; cytostome at base of deep tubular pocket; densely ciliated; common in anaerobic
habitats. Class Plagiopylea (Plagiopyla; Fig. 24M)
87b.
Body reniform, indented in middle on one side; oral groove passes around groove toward mouth; division in a cyst. Class Colpodea,
Order Colpodida, Colpodidae (Bresslaua, Tillina, Colpoda; Fig. 24N,P – S.)
LITERATURE CITED
Albright, L. J., Sherr, E. B., Sherr, B. F., Fallon, R. D. 1987. Grazing
of ciliated protozoa on free and particle-associated bacteria.
Marine Ecology Progress Series 38:125 – 129.
Allen, R. D. 1984. Paramecium phagosome membrane: from oral region to cytoproct and back again. Journal of Protozoology
31:1 – 6.
Archbold, J. H. G., Berger, J. 1985. A qualitative assessment of some
metazoan predators of Halteria grandinella, a common freshwater ciliate. Hydrobiologia 126:97 – 102.
Arndt, H. 1993. Rotifers as predators on components of the microbial food web (bacteria, heterotrophic flagellates, ciliates) — a
review. Hydrobiologia 255:193 – 204.
Balczon, J. M., Pratt, J. R. 1996. The functional response of two benthic algivorous ciliated protozoa with differing feeding strategies. Microbial Ecology 31:209 – 224.
Baldock, B. M. 1986. Peritrich ciliates epizoic on larvae of
Brachycentrus subnilus (Trichoptera): importance in relation to
the total protozoan population in streams. Hydrobiologia
131:125 – 131.
Baldock, B. M., Baker, J. H., Sleigh, M. A. 1980. Laboratory growth
rates of six species of freshwater Gymnamoebia. Oecologia
(Berlin) 47:156 – 159.
Baldock, B. M., Berger, J. 1984. Effects of low temperature on the
growth of four fresh-water amoebae (Protozoa: Gymnamoebia).
Transactions of the American Microscopical Society 103:
233 – 239.
Baldock, B. M., Sleigh, M. A. 1988. The ecology of benthic protozoa
in rivers — seasonal variation in numerical abundance in fine
sediments. Archive für Hydrobiologie 111:409 – 422.
Bearden, A. P., Schulz, T. W. 1997. Structure-activity relationships for
Pimephales and Tetrahymena: a mechanism of action approach.
Environmental Toxicology and Chemistry 16:1311 – 1317.
Beaver, J. R., Crisman, T. L. 1989. The role of ciliated protozoa in
pelagic freshwater ecosystems. Microbial Ecology 17:111 – 136.
Bell, G. 1988. Sex and death in Protozoa, Cambridge University
Press, Cambridge, MA.
90
W. D. Taylor and R. W. Sanders
Bennett, S. J., Sanders, R. W., Porter, K. G. 1990. Heterotrophic, autotrophic and mixotrophic nanoflagellates: seasonal abundances
and bacterivory in a eutrophic lake. Limnology and Oceanography 35:1821 – 1832.
Berger, J. 1979. The feeding behavior of Didinium nasutum on an
atypical prey ciliate (Colpidium campylum). Transactions of the
American Microscopical Society 141:261 – 275.
Berninger, U.-G., Finlay, B. J., Canter, H. M. 1986. The spatial distribution and ecology of zoochlorellae-bearing ciliates in a productive pond. Journal of Protozoology 33:557 – 563.
Berninger, U.-G., Caron, D. A., Sanders, R. W. 1992. Mixotrophic algae in three ice-covered lakes of the Pocono Mountains, USA.
Freshwater Biology 28:263 – 272.
Bick, H. 1972. Ciliated protozoa, An illustrated guide to the species
used as biological indicators in freshwater biology, World
Health Organization, Geneva. 198 pp.
Bird, D. F., Kalff, J. 1987. Algal phagotrophy: regulating factors and
importance relative to photosynthesis in Dinobryon (Chrysophyceae). Limnology and Oceanography 32:277 – 284.
Bockstahler, K. R., Coats, D. W. 1993. Grazing of the mixotrophic
dinoflagellate Gymnodinium sanguineum on ciliate populations
of Chesapeake Bay. Marine Biology 116:477 – 487.
Bott, T. L., Kaplan, L. A. 1989. Densities of benthic protozoa and nematodes in a Piedmont stream. Journal of the North American
Benthological Society 8:187 – 196.
Bott, T. L., Kaplan, L. A. 1990. Potential for protozoan grazing of
bacteria in streambed sediments. Journal of the North American
Benthological Society 9:336 – 345.
Bourelly, P. 1968. Les algues d’eau douce, Vol. 2. N. Boubée, Paris.
Bovee, E. C. 1985. Classes Lobosea Carpenter; Filosea Leidy; Granuloreticulosa DeSaedeleer; Order Athalamida Haeckel, in: Lee, J. J.
Hutner, S. H., Bovee, E. C. Eds. An illustrated guide to the Protozoa, Society of Protozoologists, Lawrence, Kansas pp. 158 – 252.
Bovee, E. C., Jahn, T. L. 1973. Taxonomy and phylogeny, in: Jeon,
K. W. Ed. The biology of Amoeba, Academic Press, New York,
pp. 37 – 82.
Burns, C. W., Gilbert, J. J. 1993. Predation on ciliates by freshwater
calanoid copepods: rates of predation and relative vulnerabilities of prey. Freshwater Biology 30:377 – 393.
Buskey, E. J., Stoecker, D. K. 1989. Behavioral responses of the marine tintinnid Favella sp. to phytoplankton: influence of chemical, mechanical and photic stimuli. Journal of Exerimental Marine Biology and Ecology 132:1 – 16.
Cairns, J. Jr. 1974. Protozoans (Protozoa). in: Hart C. W. Jr., Fuller,
S. L. H. Eds. Pollution ecology of freshwater invertebrates. Academic Press, New York, pp. 1 – 18.
Cairns, J. Jr., Kuhn, D. L., Plafkin, J. L. 1979. Protozoan colonization of artificial substrates, in: Wetzel, R. L. Ed. Methods and
measurements of periphyton communities: a review, Special
Technical Publication 690, American Society for Testing and
Materials, Philadelphia, Pennsylvania, pp. 34 – 57.
Cairns, J. Jr., Lanza, G. R., Parker, B. C. 1972. Pollution related
structural and functional changes in aquatic communities with
emphasis on freshwater algae and protozoa. Proceedings of the
Academy of Natural Sciences of Philadelphia 124:79 – 127.
Cairns, J. Jr., Yongue, W. H. 1977. Factors affecting the number of
species in freshwater protozoan communities, in: Cairns J. Jr.,
Eds. Aquatic microbial communities. Garland Publishing, New
York, pp. 257 – 303.
Calaway, W. T., Lackey, J. B. 1962. Waste treatment protozoa,
Florida Engineering and Industrial Experiment Station,
Gainesville, FL.
Calow, P. 1977. Conversion efficiencies in heterotrophic organisms.
Biological Reviews of the Cambridge Philosophical Society
52:385 – 409.
Canella, M. F., Rocchi-Canella, I. 1976. Biologie des Ophryoglenina.
Contributions à la connaissance des Ciliés VIII. Annali dell Universitià Università di Ferrara. (Nuova Serie) 3:suppl. 2.
Canter, H. M. 1973. A new primitive protozoan devouring centric
diatoms in the plankton. Zoological Journal of the Linnean
Society 52:63 – 83.
Carlough, L. A., Meyers, J. L. 1991. Bacterivory by sestonic protists
in a southeastern blackwater river. Limnology and Oceanography 36:873 – 883.
Caron, D. A. 1983. Technique for enumeration of heterotrophic and
phototrophic nannoplankton, using epifluorescence microscopy,
and comparison with other procedures. Applied and Environmental Microbiology 46:491 – 498.
Caron, D. A. 1987. Grazing of attached bacteria by heterotrophic
microflagellates. Microbial Ecology 13:203 – 218.
Caron, D. A., Goldman, J. C., Dennett, M. R. 1988. Experimental
demonstration of the roles of bacteria and bacterivorous protozoa in plankton nutrient cycles. Hydrobiologia 159:27 – 40.
Carrias, J.-F., Amblard, C., Bourdier, G. 1996. Protistan bacterivory
in an oligomesotrophic lake: importance of attached ciliates and
flagellates. Microbial Ecology 31:249 – 268.
Carrick, H. J., Fahnenestiel, G. L., Taylor, W. D. 1992. Growth and
production of planktonic protozoa in Lake Michigan: in situ
versus in vitro comparisons and importance to food web dynamics. Limnology and Oceanography 37:1221 – 1235.
Carter, R. F. 1968. Primary amoebic meningoencephalitis: clinical,
pathological and epidemiological features of six fatal cases.
Journal of Pathology and Bacteriology 96:1 – 25.
Cavalier-Smith, T. 1980. r- and K-tactics in the evolution of protist
developmental systems: genome size, phenotype diversifying selection, and cell cycle patterns. BioSystems 12:43 – 59.
Chapman-Andresen, C. 1973. Endocytic processes. in: Jeon, K. W.
Ed. The biology of Amoeba. Academic Press, New York, pp.
319 – 348.
Chardez, D. 1985. Protozoaires prédateurs de Thécamoebiens. Protistologica 21:187 – 194.
Cleven, E.-J. 1996. Indirectly fluorecently labelled flagellates (IFLF):
a tool to estimate the predation on free-living heterotrophic
flagellates. Journal of Plankton Research 18:429 – 442.
Cooley, N. R., Keltner, J. M. Jr., Forester, J. 1972. Mirex and
Aroclor® 1254: effect on and accumulation by Tetrahymena
pyriformis. Journal of Protozoology 19:636 – 638.
Corliss, J. O. 1959. An illustrated guide to the higher groups of the
ciliated protozoa, with a definition of terms. Journal of Protozoology 6:265 – 284.
Corliss, J. O. 1973. History, taxonomy, and evolution of species of
Tetrahymena, in Elliott, A. M. Ed. Biology of Tetrahymena,
Dowden, Hutchinson and Ross, Stroudsburg, PA, pp. 1 – 55.
Corliss, J. O. 1979. The ciliated protozoa, Characterization, classification and guide to the literature, Pergamon Press, Oxford.
Corliss, J. O., Daggett, P.-M. 1983. “Paramecium aurelia” and
“Tetrahymena pyriformis”: current status of the taxonomy and
nomenclature of these popularly known and widely used ciliates. Protistologica 19:307 – 322.
Corliss, J. O., Esser, S. C. 1974. Comments on the role of the cyst in
the life cycle and survival of free-living protozoa, Transactions
of the American Microscopical Society 93:578 – 593.
Crawford, D. W., Stoecker, D. K. 1996. Carbon content, dark respiration and mortality of the mixotrophic planktonic ciliate
Strombidium capitatum, Marine Biology 126:415 – 422.
Curds, C. R. 1963. The flocculation of suspended matter by Paramecium caudatum. Journal of General Microbiology 33:357 – 363.
Curds, C. R. 1977. Microbial interactions involving protozoa, in:
Skinner, F. A., Shewan, J. M. Eds. Aquatic Microbiology, Academic Press, New York, pp. 69 – 105.
3. Protozoa
Curds, C. R., Cockburn, A. 1970. Protozoa in biological sewagetreatment processes. II. Protozoa as indicators in the activated
sludge process. Water Research 4:237 – 249.
Curds, C. R., Cockburn, A. 1971. Continuous monoxenic culture of
Tetrahymena pyriformis. Journal of General Microbiology
66:95 – 108.
Daniels, E. W. 1973. Ultrastructure. in: Jeon, K. W. Ed., The biology
of Amoeba. Academic Press, New York, pp. 125 – 169.
Debiase, A. E., Sanders, R. W., Porter, K. G. 1990. Relative nutritional value of ciliate protozoa and algae as food for Daphnia.
Microbial Ecology 19:199 – 210.
Decamp, O., Warren, A. 1999. Bacterivory in ciliates isolated from
constructed wetlands (reed beds) used for wastewater treatment. Water Research 32:1989 – 1996
Deflandre, G. 1959. Rhizopoda and Actinopoda, in: W. T. Edmondson, editor. Freshwater Biology. Wiley, New York, pp. 232 – 264
Dhanarj, P. S., Caushal, B. R., Lal, R. 1989. Effects on uptake by the
metabolism of aldrin and phorate in a protozoan, Tetrahymena
pyriformis. Acta Protozoologica 28:157 – 163.
Dive, D. 1973. Nutrition holozoique de Colpidium campylum aux
dépens de bactéries pigmentées ou synthétisant des toxines. Protistologica 10:517 – 525.
Dodge, J. D., Gruet, C. 1987. Dinoflagellate ultrastructure and complex organelles. in: Taylor, F. J. R. Ed., The biology of Dinoflagellates. Academic Press, New York, pp. 92 – 142.
Doucet, C. M., Maly, E. J. 1990. Effects of copper on the interaction
between the predator Didinium nasutum and its prey Paramecium caudatum. Canadian Journal of Fisheries and Aquatic
Sciences 47:1122 – 1127.
Dragesco, J. 1966a. Observations sur quelques ciliés libres. Archiv
für Protistenkunde 109:155 – 206.
Dragesco, J. 1966b. Ciliés libres de thonon et sus environs. Protistologica 2:59 – 95.
Eddy, S. 1930. The freshwater armored or thecate dinoflagellates.
Transactions of the American Microscopical Society 49:
277 – 321.
Egerter, D. E., Anderson, J. R., Washburn, J. O. 1986. Dispersal of
the parasitic ciliate Lambornella clarkii: implications for ciliates
in the biological control of mosquitoes. Proceedings of the National Academy of Sciences 83:7335 – 7339.
Elliott, A. M., Ed. 1973. Biology of Tetrahymena. Dowden, Hutchinson and Ross, Stroudsburg, PA.
Evans, M. S., Sell, D. W., Beeton, A. M. 1981. Tokophrya quadripartita and Tokophrya sp. (Suctoria) associations with crustacean
zooplankton in the Great Lakes region. Transactions of the
American Microscopical Society 100:384 – 391.
Fauré-Fremiet, E. 1967. Chemical aspects of ecology, in: Florkin, M.
Scheer, B. T., Eds. Chemical ecology, Vol. 1, Protozoa. Academic Press, NewYork, pp. 21 – 54.
Fenchel, T. 1967. The ecology of the marine microbenthos. I. The
quantitative importance of ciliates as compared with metazoans
in various types of sediments. Ophelia 4:121 – 137.
Fenchel, T. 1969. The ecology of the marine microbenthos. IV. Structure and function of the benthic ecosystem, its chemical and
physical factors and the microfauna communities with special
reference to the ciliated protozoa. Ophelia 6:1 – 182.
Fenchel, T. 1974. Intrinsic rate of increase: the relationship with
body size. Oecologia (Berlin) 14:317 – 326.
Fenchel, T. 1980. Suspension feeding in ciliated protozoa: functional
response and particle size selection. Microbial Ecology 6:1 – 11.
Fenchel, T. 1982a. Ecology of heterotrophic microflagellates. I. Some
important forms and their functional morphology. Marine Ecology Progress Series 8:211 – 223.
Fenchel, T. 1982b. Ecology of heterotrophic microflagellates. II. Bioenergetics and growth. Marine Ecology Progress Series 8:225 – 231.
91
Fenchel, T. 1987. Ecology of Protozoa. Springer-Verlag, Berlin.
Fenchel, T., Perry, T., Thane, A. 1977. Anaerobiosis and symbiosis with
bacteria in free-living ciliates. Journal of Protozoology 24:154–163.
Fenchel, T., Finlay, B. J. 1991. The biology of free-living anaerobic
ciliates. European Journal of Protistology 26:201 – 215.
Fenchel, T., Estaban, G. E., Finlay, B. J. 1997. Local versus global diversity of microorganisms: cryptic diversity of ciliated protozoa.
Oikos 80:220 – 225.
Fernandez-Leborans, G., Novillo-Vajos, A. 1993. Changes in the
structure of a freshwater protozoan community subjected to cadmium. Ecotoxicology and Environmental Safety 25:271 – 279.
Finlay, B. J. 1977. The dependence of reproductive rate on cell size
and temperature in freshwater ciliated protozoa. Oecologia
(Berlin) 30:75 – 81.
Finlay, B. J. 1990. Physiological ecology of free-living protozoa. Advances in Microbial Ecology 12:1 – 35.
Finlay, B. J., Berninger, U.-G., Stewart, L. J., Hindle, R. M. Davison,
W. 1987. Some factors controlling the distribution of two ponddwelling ciliates with algal symbionts (Frontonia vernalis and
Euplotes daideleos). Journal of Protozoology 34:349 – 356.
Finlay, B. J., Corliss, J. O., Esteban, G., Fenchel, T. 1996. Biodiversity at the microbial level: the number of free-living ciliates in
the biosphere. Quarterly Review of Biology 71:221 – 237.
Finlay, B. J., Ochsenbein-Gattlen, C. 1982. Ecology of free-living
protozoa. A bibliography. Occasional paper No. 17, Freshwater
Biological Association, Ambleside, Cumbria, UK.
Finlay, B. J., Rogerson, A., Cowling, A. J. 1988. A beginners guide to
the collection, isolation, cultivation and identification of freshwater protozoa. Culture Collection of Algae and Protozoa,
Freshwater Biological Association, Ambleside, England.
Foissner, W. 1988. Taxonomic and nomenclatural revision of Sládecek’s list of ciliates (Protozoa: Ciliophora) as indicators of water
quality. Hydrobiologia 166:1 – 64.
Foissner, W., Berger, H. 1996. A user-friendly guide to the ciliates
(Protozoa: Ciliophora) commonly used by hydrobiologists as
bioindicators in rivers, lakes, and waste waters, with notes on
their ecology. Freshwater Biology 35:375 – 482.
Foissner, W., Berger, H., Schaumburg, J. 1999. Identification and
ecology of limnetic plankton ciliates. Informations berichte des
Bayer. Landesamtes für Wasserwirtschaft, Heft 3/99.
Gaines, G., Elbrächter, M. 1987. Heterotrophic nutrition, in: Taylor,
F. J. R. Ed. The biology of Dinoflagellates, Academic Press,
New York, pp. 224 – 268.
Gates, M. A. 1978. An essay on the principles of ciliate systematics.
Transactions of the American Microscopical Society 97:221 – 235
Gause, G. F. 1934. The struggle for existence. Williams and Wilkins,
Baltimore, MD.
Giese, A. C. 1973. Blepharisma. Stanford University Press, Stanford,
CA.
Gifford, D. J. 1985. Laboratory culture of marine planktonic oligotrichs (Ciliophora; Oligotrichida). Marine Ecology Progress
Series 23:257 – 267.
Gilbert, J. J. 1994. Jumping behavior in the oligotrich ciliates
Strobilidium velox and Halteria grandinella, and its significance
as a defense against rotifer predators. Microbial Ecology
27:189 – 200.
Gilbert, J. J., Jack, J. D. 1993. Rotifers and predators on small
ciliates. Hydrobiologia 255/256:247 – 253.
Goldman, J. C., Caron, D. A. 1985. Experimental studies on an
omnivorous microflagellate: implications for grazing and nutrient recycling in the marine microbial food chain. Deep-Sea
Research 32:899 – 915.
Goldman, J. C., Caron, D. A., Andersen, O. K., Dennett, M. R.
1985. Nutrient cycling in a microflagellate food chain. Marine
Ecology Progress Series 24:231 – 242.
92
W. D. Taylor and R. W. Sanders
Golini, V. I., Corliss, J. O. 1981. A note on the occurrence of the
hymenostome ciliate Tetrahymena in chironomid larvae. Transactions of the American Microscopical Society 100:89 – 93.
Goodrich, J. P., Jahn, T. L. 1943. Epizoic suctoria (Protozoa) from
turtles. Transactions of the American Microscopical Society
62:245 – 253.
Goulder, R. 1972. Grazing by the ciliated protozoon Loxodes magnus on the alga Scenedesmus in a eutrophic pond. Oecologia
(Berlin) 13:177 – 82.
Goulder, R. 1980. The ecology of two species of primitive ciliated
protozoa commonly found in standing freshwaters (Loxodes
magnus Stokes and L. striatus Penard). Hydrobiologia 72:
131 – 158.
Grell, K. G. 1973. Protozoology. Springer-Verlag, Berlin.
Grolière, C.-A. 1980. Morphologie et stomatogenèse chez deux
ciliés Scuticociliatida des genres Philasterides Kahl, 1926 et
Cyclidium O.F. Müller, 1786. Acta Protozoologica 19:
195 – 206.
Grover, J. P. 1989. Effects of Si:P supply ratio, supply variability, and
selective grazing in the plankton: an experiment with a natural
algal and protistan assemblage. Limnology and Oceanography
34:349 – 367.
Gunderson, J. H., Elwood, H., Ingold, A., Kindle, K., Sogin, M. L.
1987. Phylogenetic relationships between chlorophytes, chrysophytes, and oomycetes. Proceedings of the National Academy of
Sciences 84:5823 – 5827.
Guhl, B. E., Finlay, B. J., Schink, B. 1996. Comparison of ciliate
communities in the anoxic hypolimnia of three lakes: general
features and the influence of lake characteristics. Journal of
Plankton Research 18:335 – 353.
Gupta, R. S., Golding, G. B. 1996. The origin of the eukaryotic cell.
Trends in Biochemical Science 21:166 – 171.
Haack-bell, B., Hohenberger, R., Plattner, H. 1990. Trichocysts of
Paramecium: secretory organelles in search of their function.
European Journal of Protistology 25:289 – 305.
Hairston, N. G. 1967. Studies on the limitation of a natural population of Paramecium aurelia. Ecology 48:904 – 910.
Hairston, N. G., Kellerman, S. L. 1964. Paramecium ecology:
electromigration of field samples and observations on density.
Ecology 45:373 – 376.
Hairston, N. G., Kellerman, S. L. 1965. Competition between
varieties 2 and 3 of Paramecium aurelia: the influence of
temperature in a food-limited system. Ecology 46:134 – 139.
Hanson, E. D. 1977. The origin and early evolution of animals,
Wesleyan University Press, Middletown, CT.
Hardin, G. 1943. Flocculation of bacteria by protozoa. Nature
(London) 151:642
Harumoto, T. 1994. The role of trichocyst discharge and backward
swimming behavior of Paramecium from Dileptus margaritifer.
Journal of Eukaryotic Microbiology 41:560 – 564.
Hausmann, K. 1978. Extrusive organelles of protozoa. International
Review of Cytology 52:197 – 276.
Havens, K. E., Beaver, J. R. 1997. Consumer vs. resource control of
ciliate protozoa in a copepod-dominated lake. Archiv für
Hydrobiologie 140:491 – 511.
Henebry, M. S., Ridgeway, R. T. 1980. Epizoic ciliated protozoa of
planktonic copepods and cladocerans and their possible use as
indicators of organic water pollution. Transactions of the
American Microscopical Society 98:495 – 508.
Hewett, S. W. 1980a. Prey-dependent cell size in a protozoan predator. Journal of Protozoology 27:311 – 313.
Hewett, S. W. 1980b. The effect of prey size on the functional response and numerical response of a protozoan predator to its
prey. Ecology 61:1075 – 1081.
Hofeneder, H. 1930. Über die animalische Ernährung con Ceratium
hirudinella O.F. Müller and über die Rolle des Kernes bei dieser
Zellfunkton. Archiv für Protistenkunde 71:1 – 32.
Holwill, M. E. J. 1974. Hydrodynamic aspects of ciliary and flagellar
movement, in: Sleigh, M. A. Ed. Cilia and flagella, Academic
Press, New York, pp. 143 – 175.
Jack, J. D., Gilbert, J. J. 1997. Effects of metazoan predators on ciliates in freshwater plankton communities. Journal of Eukaryotic
Microbiology 44:194 – 199.
Jackson, K. M., Berger, J. 1985. Life history attributes of some ciliated protozoa. Transactions of the American Microscopical
Society 104:53 – 62.
Jacobson, D. M., Anderson, D. M. 1986. Thecate heterotrophic
dinoflagellates: feeding behavior and mechanisms. Journal of
Phycology 22:249 – 258.
Jahn, T. L., McKibben, W. R. 1937. A colorless euglenoid flagellate,
Khawkkinea halli n. gen., n. sp. Transactions of the American
Microscopical Society 56:48 – 54.
Jankowski, A. W. 1964a. Morphology and evolution of Ciliophora.
III. Diagnoses and phylogenesis of 53 sapropelebionts, mainly
of the order Heterotrichida. Archive für Protistenkunde
107:185 – 294.
Jankowski, A. W. 1964b. Morphology and evolution of Ciliophora.
IV. Sapropelebionts of the family Loxocephalidae fam. nova,
their taxonomy and evolutionary history. Acta Protozoologica
2:33 – 58.
Jeon, K. W., Ed. 1973. The biology of Amoeba. Academic Press,
New York.
Jeon, K. W., Ed. 1983. Intracellular symbiosis. International Review
of Cytology. Supplement 14.
Johnson, P. W., Donaghay, P. L., Small, E. B., Sieburth, J. McN:
1995. Ultrastructure and ecology of Perispira ovum (Ciliophora: Litostomatea): and aerobic, planktonic ciliate that sequesters the chloroplasts, mitochondria, and paramylon of Euglena proxima in a microoxic habitat. Journal of Eukaryotic
Microbiology 57:323 – 335.
Jones, H. L. J. 1997. A classification of mixotrophic protists based
on their behaviour. Freshwater Biology 37:35 – 43.
Jones, R. I., Rees, S. 1994. Influence of temperature and light on particle ingestion by the freshwater phytoflagellate Dinobryon.
Archiv für Hydrobiologie 132:203 – 211.
Kahl, A. 1930 – 1935. Urtiere oder Protozoa. I: Wimpertiere oder Ciliata (Infusoria), in: Dahl, F. Ed. Die Tierwelt Deutschlands 18
(1930), 21 (1931), 25 (1932), 30 (1935). Fischer-Verlag, Jena,
pp. 1 – 886.
Kankaala, P., Eloranta, P. 1987. Epizooic ciliates (Vorticella sp.) compete for food with their host Daphnia longispina in a small
polyhumic lake. Oecologia (Berlin) 73:203 – 206.
Kemp, P. F., Sherr, B. F., Sherr, E. B., Cole, J. J. 1993. Handbook of
methods in aquatic microbial ecology, Lewis Publishers, Boca
Raton.
Kent, W. S. 1881 – 1882. A manual of the infusoria, Vol. 2, David
Bogue, London.
Kentouri, M., Divanah, P. 1986. Sur l’mportance des ciliés pélagiques
dans l’alimentation des stades larvaires de poissons. (On the importance of pelagic ciliates in the feeding of larval stages of
fish.) Année Biologique 25:307 – 318.
Kerr, S. J. 1983. Colonization of blue-green algae by Vorticella
(Ciliata, Peritrichida). Transactions of the American Microscopical Society 102:38 – 47.
Kornienko, G. S. 1971. The role of infusoria in the feeding of larvae
of herbivorous fish. Journal of Ichthyology 11:241 – 246.
Kudo, R. R. 1966. Protozoology, 5th ed., C. C. Thomas, Springfield,
IL.
3. Protozoa
Kusch, J. 1993. Induction of defense morphological changes in ciliates. Oecologia (Berlin) 94:571 – 575.
Kusuoka, Y., Watanabe, Y. 1989. Distinction of emigration by
telotroch formation and death by predation in peritrich ciliates:
SEM observations on the remaining stalk ends. FEMS Microbiology Ecology 62:7 – 12.
Lackey, J. B. 1959. Zooflagellates, in: Edmondson, W. T. Ed., Freshwater Biology. Wiley, New York, pp. 190 – 231.
Lal, S., Saxena, B. M., Lal. R. 1987. Uptake, metabolism and effects
of DDT, Fenthrothion and chlorphyrifos on Tetrahymena pyriformis. Pesticide Science 21:181 – 191.
Landis, W. G. 1981. The ecology of the killer trait, and interactions
of five species of the Paramecium aurelia complex inhabiting
the littoral zone. Canadian Journal of Zoology 59:1734 – 1743.
Lawler, S. P. 1993. Direct and indirect effects in microcosm communities of protists. Oecologia. 93:184 – 190.
Laybourn-Parry, J. 1984. A functional biology of free-living Protozoa. University of California Press, Berkley, CA.
Laybourn-Parry, J. 1992. Protozoan plankton ecology. Chapman &
Hall, London/New York.
Laybourn-Parry, J., Bayliss, P., Ellis-Evans, J. C. 1995. The dynamics
of heterotrophic nanoflagellates and bacterioplankton in a large
ultra-oligotrophic Antarctic lake. Journal of Plankton Research
17:1835 – 1850.
Leadbeater, B. S. C., Morton, C. 1974. A microscopical study of a
marine species of Codosiga James-Clark (Choanoflagellata)
with special reference to the ingestion of bacteria. Biological
Journal of the Linnean Society 6:337 – 347.
Lee, J. J., Corliss, J. O. 1985. Symposium on “Symbiosis in Protozoa”:
introductory remarks. Journal of Protozoology 32:371 – 372.
Lee, J. J., Small, E. B., Lynn, D. H., Bovee, E. C. 1985. Some techniques for collecting, cultivating, and observing protozoa, in
Lee, J. J., Hutner, S. H., Bovee, E. C. Eds. An illustrated guide
to the Protozoa, Society of Protozoologists, Lawrence, KS,
pp. 1 – 7.
Lee, C. C., Fenchel, T. 1972. Studies on ciliates associated with sea
ice from Antarctica. II. Temperature responses and tolerances in
ciliates from Antarctic, temperate and tropical habitats. Archive
für Protistenkunde 114:237 – 244.
Leedale, G. F. 1985. Order 3. Euglenida Bütschli, 1984, in: Lee, J. J.,
Hutner, S. H., Bovee, E. C., Eds., An illustrated guide to the
Protozoa. Society of Protozoologists. Lawrence, KS, pp. 41 – 54.
Lemmerman, E. 1914. Protomastiginae, in: Pascher, A. Ed. Die süsswasser-flora deutschlands, österreichs und der schweiz, I. Flagellatae I. Fischer, Jena, pp. 30 – 121.
Lim, E. L. 1996. Molecular identification of nanoplanktonic protists
based on small subunit ribosomal RNA gene sequences for ecological studies. Journal of Eukaryotic Microbiology 43:
101 – 106.
Luckinbill, L. S. 1979. Selection and the r/K continuum in experimental populations of protozoa. American Naturalist 113:
427 – 437.
Luckinbill, L. S., Fenton, M. M. 1978. Regulation and environmental variability in experimental populations of protozoa. Ecology
59:1271 – 1276.
Lynn, D. H. 1975. The life cycle of the histophagous ciliate Tetrahymena corlissi Thompson, 1955. Journal of Protozoology
22:188 – 195.
Lynn, 1976. Comparative ultrastructure and systematics of the
Colpodida. Structural conservatism hypothesis and a description of Colpoda steini Maupas. Journal of Protozoology
23:302 – 314.
Lynn, 1977. Comparative ultrastructure and systematics of the
Colpodida. Fine structure specializations associated with large
93
body size in Tillina magna Gruber, 1880. Protistologica
12:629 – 648.
Lynn, D. H. 1981. The organization and evolution of the microtubular organelles in ciliated protozoa. Biological Reviews of the
Cambridge Philosophical Society 56:243 – 292.
Lynn, D. H., Corliss, J. O. 1990. Phylum Ciliophora (Ciliates), in:
Harrison, F. W., Corliss, J. O., Eds., Microscopic anatomy of
invertebrates, Vol. 1: The Protozoa. Wiley-Liss, New York,
pp. 331 – 465.
Lynn, D. H., Small, E. B. 1997. A revised classification of the
phylum Ciliophora Doflein, 1901. Rev. Soc. Mex. Hist. Nat.
47:65 – 78.
Maguire, B. Jr. 1963a. The passive dispersal of small aquatic organisms and their colonization of isolated bodies of water. Ecological Monographs 33:161 – 185.
Maguire, B. Jr. 1963b. The exclusion of Colpoda (Ciliata) from
superficially favorable habitats. Ecology 44:781 – 784.
Maguire, B. Jr. 1977. Community structure of protozoans and algae
with particular emphasis on recently colonized bodies of water,
in: Cairns, J. Jr., Ed. Aquatic microbial communities, Garland
Publishing, New York, pp. 358 – 397.
Margulis, L. 1981. Symbiosis in cell evolution. W.H. Freeman, San
Francisco, CA.
Margulis, L., Corliss, J. O., Melkonian, M., Chapman, D. J., Eds.
1989. Handbook of Protoctista, Jones and Bartlett Publishers,
Boston.
Margulis, L., Schwartz, K. V. 1988. Five kingdoms: an illustrated
guide to life on earth. 2nd ed., Freeman, W. H. San Francisco,
CA.
Margulis, L., Sagan, D. 1986. Origins of sex, Yale University Press,
New Haven.
Massana, R., Pedrós-Alió, C. 1994. Role of anaerobic ciliates in
planktonic food webs: abundance, feeding and impact on bacteria in the field. Applied and Environmental Microbiology
60:1325 – 1334.
Mast, S. O., Hahnert, W. F. 1935. Feeding, digestion, and starvation
in Amoeba proteus Leidy. Physiological Zoology 8:255 – 272.
Matthes, D. 1954. Suktorienstudien. VI. Die gattung Heliophrya
Saedeleer and Tellier 1929. Archiv für Protistenkunde 100:
143 – 152.
McCormick, P. V., Cairns, J. Jr. 1991. Effects of micrometazoa on the
protistan assemblage of a littoral food web. Freshwater Biology
26:111 – 119.
McManus, G. B., Fuhrman, J. A. 1986. Bacterivory in seawater studied with the use of inert fluorescent particles. Limnology and
Oceanography 31:420 – 426.
Meisterfeld, R. 1991. Vertical distribution of Difflugia hydrostatica
(Protozoa, Rhizopoda). Internationale Vereinigung für
Theoretische und Angewandte Limnologie Verhandlungen
24:2726 – 2728.
Montagnes, D. J. S., Lynn, D. H. 1987a. A quantitative protargol
stain (QPS) for ciliates: a method description and test of its
quantitative nature. Marine Microbial Food Webs 2:83 – 93.
Montagnes, D. J. S., Lynn, D. H. 1987b. Agar embedding on
cellulose filters: an improved method of mounting protists for
protargol and Chatton-Lwoff staining. Transactions of the
American Microscopical Society 106:183 – 186.
Müller, H., Geller, W. 1993. Maximum growth rates of aquatic
ciliated protozoa: the dependence on body size and temperature
reconsidered. Archiv für Hydrobiologie 126:315 – 327.
Nagata, T. 1988. The microflagellate-picoplankton food linkage in
the water column of Lake Biwa. Limnology and Oceanography
33:504 – 517.
Nilsson, J. R. 1976. Phagotrophy in Tetrahymena. in Levandowsky, M.
94
W. D. Taylor and R. W. Sanders
and Hutner, S. H., Eds. Biochemistry and Physiology of Protozoa.
Vol. 2., Academic Press, New York, pp. 339 – 379.
Nilsson, J. R. 1989. Tetrahymena in cytotoxicology: with special reference to effects of heavy metals and selected drugs. European
Journal of Protistology 25:2 – 25.
Nilsson, J. R. 1987. Structural aspects of digestion of Escherichia coli
in Tetrahymena. Journal of Protozoology 34:1 – 6.
Nisbet, B. 1984. Nutrition and feeding strategies in Protozoa. Croom
Helm, London.
Noland, L. E. 1925. The factors affecting the distribution of fresh
water ciliates. Ecology 6:437 – 452.
Noland, L. E. 1959. Ciliophora, in: Edmondson, W. T. Ed., Freshwater Biology, Wiley, New York, pp. 265 – 297.
Noland, L. E., Goldjics, M. 1967. Ecology of free-living Protozoa, in:
Chen, T.-T. Ed. Research in protozoology. Vol. 2, Pegamon, Oxford, pp. 215 – 266.
Nyberg, D. 1974. Breeding systems and resistance to environmental
stress in ciliates. Evolution 28:367 – 380.
Nyberg, D., Bishop, P. 1983. High levels of phenotypic variability of
metal and temperature tolerance in Paramecium. Evolution
37:341 – 357.
Nygaard, K., Borsheim, K. Y., Thingstad, T. F. 1988. Grazing rates
on bacteria by marine heterotrophic microflagellates compared
to uptake rates of bacteria-sized monodisperse fluorescent latex
beads. Marine Ecology Progress Series 44:159 – 165.
Ogden, C. G. 1991. Gas vacuoles and flotation in the testate amoeba
Arcella discoides. Journal of Protozoology 38:217 – 220.
Pace, M. L., Orcutt, J. D. Jr. 1981. The relative importance of protozoans, rotifers, and crustaceans in a freshwater zooplankton
community. Limnology and Oceanography 26:822 – 830.
Page, F. C. 1976. An illustrated key to freshwater and soil amoebae,
with notes on cultivation and ecology. Scientific Publication
Number 34, Freshwater Biological Association, Ambleside,
Cumbria, England.
Page, F. C. 1977. The genus Thecamoeba (Protozoa, Gymnamoebia).
Species distinctions, locomotive morphology, and protozoan
prey. Journal of Natural History 11:25 – 63.
Page, F. C. 1987. The classification of ‘naked’ amoebae (Phylum
Rhizopoda). Archive für Protistenkunde 133:199 – 217.
Page, F. C. 1988. A new key to freshwater and soil Gymnamoebae,
Freshwater Biological Association, Ambleside, England.
Pascher, A. 1913. Chrysomonadidae, in: Pascher, A. Ed., Die
süsswasser-flora deutschlands, österreichs und der schweiz, I.
Flagellatae II. Fischer, Jena, pp. 7 – 95.
Patterson, D. J. 1980. Contractile vacuoles and associated structures:
their organization and function. Biological Reviews of the Cambridge Philosophical Society 55:1 – 46.
Patterson, D. J. 1993. The current status of the free-living
heterotrophic flagellates. Journal of Eukaryotic Microbiology
40:606 – 608.
Peck, R. K. 1985. Feeding behavior in the ciliate Pseudomicrothorax
dubius is a series of morphologically and physiologically distinct events. Journal of Protozoology 32:492 – 501.
Picken, L. E. R. 1937. The structure of some protozoan communities.
Journal of Ecology 25:368 – 384.
Pomeroy, L. R. 1974. The ocean’s foodweb: a changing paradigm.
BioScience 24:499 – 504.
Porter, K. G. 1988. Phagotrophic flagellates in microbial food webs.
Hydrobiologia 159:89 – 97.
Porter, K. G., Sherr, E. B., Sherr, B. F., Pace, M. C., Sanders, R. S.
1985. Protozoa in planktonic food webs. Journal of Protozoology 32:409 – 415.
Pratt, J. R., Cairns, J. Jr. 1985. Functional groups in the protozoa:
roles in differing ecosystems. Journal of Protozoology 32:
415 – 423.
Pratt, J. R., Rosen, B. H. 1983. Associations of species of Vorticella
(Peritrichida) and planktonic algae. Transactions of the American Microscopical Society 102:48 – 54.
Rainier, H. 1968. Heliozoa, in: Dahl, F. Ed. Die tierwelt deutschlands, Vol. 56, Fischer-Verlag, Jena, pp. 3 – 174.
Rapport, D., Berger, J., Reid, D. B. 1972. Determination of food preference of Stentor coeruleus. Biological Bulletin 142:103 – 109.
Rasmussen, L. 1976. Nutrient uptake in Tetrahymena. Carlsberg
Research Communications 41:143 – 167.
Ricci, N. 1989. Microhabitats of ciliates: specific adaptations to
different substrates. Limnology and Oceanography 34:
1089 – 1097.
Ricketts, T. R. 1972. The interaction of particulate material and dissolved foodstuffs in food uptake by Tetrahymena pyriformis.
Archive für Mikrobiologie 81:344 – 349.
Riemann, B., Christoffersen, K. 1993. Microbial trophodynamics in
temperate lakes. Marine Microbial Food Webs 7:69 – 100.
Rogerson, A., Berger, J. 1981. Effect of crude oil and petroleumdegrading microorganisms on the growth of freshwater and soil
protozoa. Journal of General Microbiology 124:53 – 59.
Rogerson, A., Finlay, B. J., Berninger, U.-G. 1989. Sequestered
chloroplasts in the freshwater ciliate Strombidium viride (Ciliophora: Oligotrichina). Transactions of the American Microscopical Society 108:117 – 126.
Rogerson, A., Shiu, W. Y., Huang, G. L., Mackay, D., Berger, J.
1983. Determination and interpretation of hydrocarbon toxicity
to ciliate protozoa. Aquatic Toxicology 3:215 – 228.
Rudzinska, M. A. 1965. The fine structure and function of the tentacles of Tokophrya infusionum and ultrastructural changes in
food vacuoles during digestion. Journal of Cell Biology 25:
459 – 477.
Ruthven, J. A., Cairns, J. Jr. 1973. Response of fresh-water protozoan artificial communities to metals. Journal of Protozoology
20:127 – 135.
Sambanis, A., Fredrickson, A. G. 1988. Persistence of bacteria in the
presence of viable, nonencysting bacterivorous ciliates. Microbial Ecology 16:197 – 212.
Sanders, R. W. 1988. Feeding by Cyclidium sp. (Ciliophora, Scuticociliatida) on particles of different sizes and surface properties.
Bulletin of Marine Science 43:446 – 457.
Sanders, R. W. 1991. Mixotrophic protists in marine and freshwater
ecosystems. Journal of Protozoology 38:76 – 80.
Sanders, R. W., Porter, K. G. 1988. Phagotrophic flagellates.
Advances in Microbial Ecology 10:167 – 192.
Sanders, R. W., Porter, K. G., Bennett, S. J., DeBiase, A. E. 1989.
Seasonal patterns of bacterivory by flagellates, ciliates, rotifers
and cladocerans in a freshwater planktonic community. Limnology and Oceanography 34:673 – 687.
Sanders, R. W., Porter, K. G., Caron, D. A. 1990. Relationship
between phototrophy and phagotrophy in the mixotrophic
chrysophyte Poterioochromonas mahlhamensis. Microbial
Ecology 19:97 – 109.
Sanders, R. W., Leeper, D. A., King, C. H., Porter, K. G. 1994. Food
web structure and zooplankton grazing impacts on photosynthetic and heterotrophic nanoplankton. Hydrobiologia 288:
167 – 181.
Sanders, R. W., Wickham, S. A. 1993. Planktonic protozoa and
metazoa: predation, food quality and population control.
Marine Microbial Food Webs 7:197 – 223.
Schönborn, W. 1977. Production studies on Protozoa. Oecologia
(Berlin) 27:171 – 184
Schuster, F. L. 1979. Small amebas and ameboflagellates. in
Levandowsky M., Hutner, S. H. Eds. Biochemistry and physiology of Protozoa, Vol. 1, Academic Press, New York, pp.
215 – 285.
3. Protozoa
Scott, J. M. 1985. The feeding rates and efficiencies of a marine
ciliate, Strombidium sp., grown under chemostat steady-state
conditions. Journal of Experimental Marine Biology and Ecology 90:81 – 95.
Scott, B. F., Wade, P. J., Taylor, W. D. 1984. Impact of oil and oildispersant mixtures on the fauna of freshwater ponds. The
Science of the Total Environment 35:191 – 206.
Shawhan, F. M., Jahn, T. L. 1947. A survey of the genus
Petalomonas Stein (Protozoa: Euglenida). Transactions of the
American Microscopical Society 66:182 – 189.
Sherr, E. B., Sherr, B. F., Albright, L. J. 1987. Bacteria: Link or sink?
Science 235:88 – 89.
Shuter, B. J., Thomas, J. E., Taylor, W. D., Zimmerman, A. M. 1983.
Phenotypic correlates of genomic DNA content in unicellular
eukaryotes and other cells. American Naturalist 122:26 – 44.
Simek, K., Straskrabová, V. 1992. Bacterioplankton production and
protozoan bacterivory in a mesotrophic reservoir. Journal of
Plankton Research 14:773 – 787.
Simek, K., Chrzanowski, T. H. 1992. Direct and indirect evidence
of size-selective grazing on pelagic bacteria by freshwater
nanoflagellates. Applied and Environmental Microbiology
58:3715 – 3720.
Simek, K., Vrba, J., Hartman, P. 1994. Size-selective feeding by
Cyclidium sp. on bacterioplankton and various sizes of cultured
bacteria. FEMS Microbial Ecology 14:157 – 168.
Singh, B. N. 1975. Pathogenic and nonpathogenic amoebae,
MacMillan, London.
Skibbe, O. 1994. An improved quantitative protargol stain for ciliates and other planktonic protists, Archiv für Hydrobiologie
130:339 – 348.
Slabbert, J. L., Morgan, W. S. G. 1982. A bioassay technique using
Tetrahymena pyriformis for the rapid assessment of toxicants in
water. Water Research 16:517 – 523.
Sládecek, V. 1973. System of water quality from the biological point
of view. Ergebnisse der Limnologie 7:1 – 218.
Sleigh, M. A., Blake, J. R. 1977. Methods of ciliary propulsion and
their size limitations, in: Pedley, T. J. Ed. Scale effects in animal
locomotion. Academic press, New York, pp. 243 – 256.
Sleigh, M. A, Baldock, B. A., Baker., J. H. 1993. Protozoan communities in chalk streams. Hydrobiologia 248:53 – 64.
Small, E. B., Lynn, D. H. 1985. Phylum Ciliophora Doflein, 1901. in:
Lee, J. J., Hutner, S. H., Bovee, E. C., Eds., An illustrated guide
to the Protozoa, Society of Protozoologists. Lawrence, KS,
pp. 393 – 575.
Smith, G. M. 1950. The fresh-water algae of the United States,
2nd ed., McGraw-Hill, New York.
Sogin, M. L., Elwood, H. J. 1986. Primary structure of the Paramecium aurelia small-subunit rRNA coding region: phylogenetic
relationships within the Ciliophora. Journal of Molecular
Evolution 23:53 – 60.
Sogin, M. L., Gunderson, J. H., Elwood, H. J., Olonso, R. A.,
Peattie, D. A. 1989. Phylogenetic meaning of the Kingdom
concept: an unusual ribosomal RNA from Giardia lamblia. Science 243:75 – 77.
Sogin, M. L., Silberman, J. D., Hinkle, G., Morrison, H. D. 1996.
Problems with molecular diversity in the Eukarya. in Roberts,
D. McC., Sharp, P., Alderson, G., Collins, M. Eds., Evolution
of microbial life. Cambridge University Press, Cambridge,
pp. 167 – 184.
Sommaruga, R., Oberleiter, A., Psenner, R. 1996. Effect of UV
radiation on the bacterivory of a heterotrophic nanoflagellate.
Applied and Environmental Microbiology 62:4395 – 4400
Sommaruga, R., Psenner, R. 1993. Nanociliates of the order
Prostomatida: their relevance in the microbial food web of a
mesotrophic lake. Aquatic Sciences 55:179 – 187.
95
Sonneborn, T. M. 1975. The Paramecium aurelia complex of fourteen sibling species. Transactions of the American Microscopical Society 94:155 – 178.
Spoon, D. M., Burbanck, W. D. 1967. A new method for collecting
sessile ciliates in plastic petri dishes with tight-fitting lids. Journal of Protozoology 14:735 – 744.
Starink, M., Krylova, I. N., Bär-Gilissen, M.-J., Bak, R. P. M.,
Cappenberg, T. E. 1994. Rates of benthic protozoan grazing on
free and attached sediment bacteria measured with fluorescently
stained sediment. Applied and Environmental Microbiology
60:2259 – 2264.
Stoecker, D. K., Capuzzo, J. M. 1990. Predation on protozoa: its
importance to zooplankton. Journal of Plankton Research
12:891 – 908.
Stoecker, D. K., Sunda, W. G., Davis, L. H. 1986. Effects of
copper and zinc on two planktonic ciliates. Marine Biology
92:21 – 29.
Stoecker, D. K., Silver, M. W., Michaels, A. E., Davis, L. H. 1989.
Enslavement of algal chloroplasts by four Strombidium spp.
(Ciliophora, Oligotrichida). Marine Microbial Food Webs
3:79 – 100.
Stout, J. D. 1980. The role of protozoa in nutrient cycling and energy
flow. Advances in Microbial Ecology 4:1 – 50.
Suttle, C. A., Chan, A. M., Taylor, W. D., Harrison, P. J. 1986. Grazing of planktonic diatoms by microflagellates. Journal of Plankton Research 8:393 – 398.
Taylor, F. J. R. 1987. Some eco-evolutionary aspects of intracellular
symbiosis, in: Jeon, K. W. Ed., Intracellular symbiosis. International Review of Cytology. Suppliment 14, pp. 1 – 28.
Taylor, W. D. 1978a. Growth responses of ciliate protozoa to the
abundance of their bacterial prey. Microbial Ecology 4:
207 – 214.
Taylor, W. D. 1978b. Maximum growth rate, size and commonness
in a community of bacterivorous ciliates. Oecologia (Berlin)
36:263 – 272.
Taylor, W. D. 1981. Temporal heterogeneity and the ecology of lotic
ciliates. in: Locke, M. A. Williams, D. D. Eds., Perspectives in
running water ecology, Plenum, New York, pp. 209 – 224.
Taylor, W. D. 1983a. A comparative study of the sessile ciliates of
several streams, Hydrobiologia 98:125 – 133.
Taylor, W. D. 1983b. Rate of population increase and mortality for
sessile ciliates in a stream riffle. Canadian Journal of Zoology
61:2023 – 2028.
Taylor, W. D. 1986. The effect of grazing by a ciliated protozoan on
phosphorus limitation in batch culture. Journal of Protozoology
33:47 – 52.
Taylor, W. D., Berger, J. 1976. Growth responses of cohabiting ciliate
protozoa to various prey bacteria. Canadian Journal of Zoology
54:1111 – 1114.
Taylor, W. D., Heynen, M. L. 1987. Seasonal and vertical distribution of Ciliophora in Lake Ontario. Canadian Journal of
Fisheries and Aquatic Sciences 44:2185 – 2191.
Taylor, W. D., Johannsson, O. 1991. A comparison of estimates of
productivity and consumption by zooplankton for planktonic
ciliates in Lake Ontario. Journal of Plankton Research
13:363 – 372.
Taylor, W. D., Lean, D. R. S. 1991. Phosphorus pool sizes and fluxes
in the epilimnion of a mesotrophic lake. Canadian Journal of
Fisheries and Aquatic Sciences 48:1293 – 1301.
Taylor, W. D., Shuter, B. J. 1981. Body size, genome size, and intrinsic rate of increase in ciliated protozoa. The American Naturalist 118:160 – 172.
Thompson, A. S., Rhodes, J. C., Pettman, I. 1988. Catalogue of
strains. Culture collection of Algae and Protozoa, Freshwater
Biological Association, Ambleside, Cumbria, England.
96
W. D. Taylor and R. W. Sanders
Tucker, J. B. 1968. Fine structure and function of the pharyngeal
basket in the ciliate Nassula. Journal of Cell Science 3:
493 – 514.
Tyndall, R. L., Ironside, K. S., Metler, P. L., Tan, E. L., Hazen, T. C.,
Fliermanns, C. B. 1989. Effect of thermal additions on the density and distribution of thermophilic amoebae and pathogenic
Naegleria fowleri in a newly created cooling lake. Applied and
Environmental Microbiology 55:722 – 732.
Vickerman, K. 1976. The diversity of kinetoplastid flagellates, in:
Lumsden, W. H. R., Evans, D. A. Eds., Biology of Kinetoplastida. Academic Press, New York, pp. 1 – 34.
Wagener, S., Schulz, S., Hanselmann, K. 1990. Abundance and distribution of anaerobic protozoa and their contribution to methane
production in Lake Cadagno (Switzerland). FEMS Microbial
Ecology 74:39 – 48.
Wang, S.-J., Heath, R. T. 1997. The distribution of protozoa across
a trophic gradient, factors controlling their abundance and
importance in the plankton food web. Journal of Plankton
Research 19:491 – 518.
Washburn, J. O., Gross, M. E., Mercer, D. R., Mercer, J. R. 1988.
Predator-induced trophic shift of a free-living ciliates: parasitism
of mosquito larvae by their prey. Science 240:1193 – 1195.
Weisse, T. 1991. The annual cycle of heterotrophic freshwater
nanoflagellates: role of bottom-up versus top-down control.
Journal of Plankton Research 13:167 – 185.
Weisse, T., Müller, H., Pinto-Coelho, R. M., Schweizer, A., Springmann, D., Baldringer, G. 1990. Response of the microbial loop
to the phytoplankton spring bloom in a large prealpine lake.
Limnol. Oceanogr. 35:781 – 794.
Wessenberg, H., Antipa, G. 1970. Capture and digestion of Paramecium by Didinium nasutum. Journal of Protozoology 17:
250 – 270.
Wickham, S. A. 1995a. Cyclops predation on ciliates: species-specific
differences and functional responses. Journal of Plankton Research 17:1633 – 1646.
Wickham, S. A. 1995b. Trophic relationships between cyclopoid
copepods and ciliated protists: complex interactions link the microbial and classic food webs. Limnology and Oceanography
40:1173 – 1181.
Wickham, S. A., Gilbert, J. J. 1993. The comparative importance of
competition and predation by Daphnia on ciliated protozoa.
Archive für Hydrobiologie 126:289 – 314.
Wicklow, B. J. 1997. Signal-induced defensive phenotypic changes in
ciliated protists: morphological and ecological implications
for predator and prey. Journal of Eukaryotic Microbiology
44:176 – 188.
Wichterman, R. 1986. The biology of Paramecium, 2nd ed., Plenum
Press, New York.
Wolfe, J. 1988. Analysis of Tetrahymena mucocyst material with
lectins and Alcian blue. Journal of Protozoology 35:46 – 51.
4
PORIFERA1
Thomas M. Frost†
Henry M. Reiswig
Redpath Museum and
Department of Biology
McGill University
Montreal, Quebec H3A 2K6
I. Introduction
II. Structure and Physiology
A. External Morphology
B. Internal Anatomy and Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
I. INTRODUCTION
Sponges in fresh water? Many people react this
way when they are told that there might be sponges in
a lake or stream. Most people have seen sponges from
the ocean but are unaware that they also occur in
freshwater. Yet sponges are common and sometimes
abundant inhabitants of a wide variety of freshwater
habitats. In some situations they comprise a major
component of the benthic fauna and may play important roles in ecosystem processes in freshwater.
Sponges are the simplest of the multicellular
phyla. They lack organs and tissues are their highest
level of organization. Specialized cells accomplish
many basic biological functions in sponges. Despite
their simplicity, however, sponges display a variety of
1
The preparation of this chapter was supported by several grants
from the National Science Foundation. We thank Janet Blair, Joan
Elias, Susan Knight, and Yolanda Lukaziewski for their assistance
in its preparation.
†
It is with profound regret that we learned in the late summer of
2000 of the death of Tom Frost who gave his life saving another
person from drowning. He will be missed by his many friends and
colleagues.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
Anthony Ricciardi
Department of Biology
Dalhousie University
Halifax, Nova Scotia B3H 4J1
IV. Collecting, Rearing, and Preparing Sponges
for Identification
V. Classification
A. Freshwater Sponge Species of Canada
and the United States
B. Key to the Freshwater Sponges in
Canada and the United States
Literature Cited
elegant adaptations to freshwater habitats including a
strong capacity for osmoregulation, complex life
cycles, a capability to feed selectively on a broad
range of particulate resources, and, in many species,
an intimate association with symbiotic algae. This
chapter will introduce you to the structure, function,
and diversity of freshwater sponges. It will emphasize
their taxonomy and their basic ecology. For those
whose interests are whetted by this introduction,
Bergquist (1978) and Simpson (1984) provide detailed
treatments of the biology of the phylum Porifera in
general.
II. STRUCTURE AND PHYSIOLOGY
The simplicity of a sponge’s external features
contrasts sharply with the complexity of its internal
structure and function. Viewed macroscopically, freshwater sponges exhibit a limited range of non-distinct
body forms, but a microscopic examination reveals a
variety of internal features. This contrast derives from
the basic organization of sponges in which tissues
rather than organs represent the highest level of
97
98
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
FIGURE 1
Gemmulated colonies of Spongilla lacustris and Duosclera mackayi exhibiting an encrusted
growth form. The distinct zone between the two species illustrates the typical nonmerging front that occurs
when two different freshwater species grow into contact with each other on a substratum.
morphological complexity. In fact, specialized cells operating independently or in association with related
cells accomplish such primary functions as food gathering, digestion, and reproduction. An appreciation of
the utility of sponge structure, therefore, is possible
only by including microscopic examinations. Likewise,
it is difficult to discuss the physiological functions of
sponges independently of their microscopic structure
and these topics are presented jointly, in a general
overview.
There has been substantial debate as to whether
sponges should be considered as colonies or individuals (Hartman and Reiswig, 1973; Simpson, 1973).
This reflects the continuing difficulties that ecologists,
evolutionary biologists, and zoologists in general encounter in deciding on what an individual is (Santelices, 1999). From an ecological perspective, it is
most important to consider that sponges are much
more like colonies, such as corals or bryozoans, in
terms of their ecological function than they are like
individual arthropods or vertebrates. Overall, however, concepts of colony or individual are not really
appropriate for sponges, because of their level of biological organization.
A. External Morphology
Freshwater sponges display a variety of morphologies that range from encrusting (Fig. 1), to rounded
(Fig. 2) and finger-like (Fig. 3) growth forms. Because
form can vary substantially within a species (Penney
and Racek, 1968), external morphology is of very
limited use in sponge taxonomy. Variation in growth
form is influenced by environmental conditions such as
water movement (Jewell, 1935; Palumbi, 1984) and the
availability of light (Frost and Williamson, 1980). Differences can be particularly dramatic between lentic
and lotic habitats (Neidhoefer, 1940).
The surface structure of sponges varies from flat
(Fig. 1) to strongly convoluted (Fig. 2). Although there
are some species-associated differences in surface characteristics, this feature, too, is strongly influenced by
habitat conditions. Finer-scale characteristics of external sponge anatomy are related to internal features.
4. Porifera
FIGURE 2 A colony of Ephydatia muelleri exhibiting a rounded growth form encrusted on a branch from a
fallen tree (from Neidhoefer, 1940).
FIGURE 3 A colony of the common freshwater sponge Spongilla lacustris exhibiting a characteristic,
fingerlike growth form.
99
100
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
Examples, discussed below, include the incurrent and
excurrent portions of the feeding canals and extensions
of the skeletal system.
B. Internal Anatomy and Physiology
The basic organization of a sponge consists of a
surface epithelium, composed of pinacocytes, surrounding an organic matrix, termed the mesohyl,
which contains a broad variety of specialized cells.
Cells within the mesohyl interact with the epidermal
cells to accomplish basic functions including the processing of water for food uptake and gas exchange, digestion, structural support, and reproduction. Some
sponge cells are highly plastic in their behavior and can
shift their function with changing environmental conditions. De Vos et al. (1991) provided a detailed atlas of
the overall general morphology of sponges illustrating
most cell types with scanning and transmission electron
micrographs.
All sponge cells, including those in the epidermis
and in the mesohyl, appear to be involved in osmoregulation (Weissenfels, 1975). This capacity has been critical in allowing the invasion of freshwater by sponges
from their original marine habitats.
FIGURE 4
1. Water Processing System
A sponge can be considered, essentially, as a series
of progressively finer filters and a mechanism that
circulates water through them. The sponge filters are
capable of removing particles ranging from large algae
to bacterial cells less than 1 m in diameter and, possibly, to colloidal organic matter (Frost, 1987). In addition, the movement of water through the canal system,
with its extensive surface area, fosters the transport of
gases and excreted materials. The main features of the
sponge water-processing system (Figs. 4 and 5) have
been identified through the efforts of numerous investigators (reviewed in Frost, 1976b, 1987) with several
papers by Weissenfels and his co-workers (Weissenfels,
1975, 1976, 1980, 198l, 1982,1992; Langenbruch and
Weissenfels, 1987) providing particularly important details on structure. Although there are some subtle differences in structures among species, the summary below provides a general overview of the structures
associated with sponge water processing.
Water first enters a sponge through a large number
of 50-m diameter openings termed ostia (single: ostium) that are spread across its surface epithelium.
Passing through the surface, inflowing water next enters a large subdermal cavity. From there, water flows
A schematic diagram indicating the primary components of the sponge feeding canal
system. The diagram is not drawn to scale, but Figure 5 provides an indication of the size and
arrangement of the feeding system in an actual sponge. Water flow patterns are indicated by arrows.
4. Porifera
101
FIGURE 5
A scanning electron micrograph of a cross section of the freshwater sponge
Ephydatia fluviatilis indicating the form, size, and arrangement of the feeding canal system.
Illustrated are a sudbdermal cavity (SD), an incurrent canal (IC) with lateral branches into the
sponge (i), and excurrent canal (EC) and its lateral branches (e), an atrium (At), and an osculum
(Os). Arrows indicate the flow path of water and numerous choanocyte chambers are visible
throughout the sponge. Magnification, approximately 100 (from Weissenfels, 1982).
into incurrent canals which branch repeatedly into
canals of progressively narrower diameters, until they
reach a choanocyte chamber (Fig. 4).
Choanocyte chambers are the keystones of the
sponge feeding system. Each chamber consists of numerous choanocytes that are characterized by the presence of a flagellum and a collar of microvilli. Beating
by the flagella within these chambers is primarily responsible for setting up the flow of water through the
sponge. Water flow is also facilitated by a sponge’s basic hydrodynamic structure. Vogel (1974) has demonstrated that water will actually flow passively, albeit at
a reduced rate, through the feeding canals of killed
sponges because of this structure. Choanocytes also
serve as the final filters in the feeding system. The microvilli that make up the collars of the choanocytes are
spaced to form openings less than 0.1 m in diameter.
Once water has passed the choanocyte chambers, it
enters the excurrent portion of the water processing
system. Exiting the choanocyte chambers, the canals
join together in an anastomosing fashion forming
larger and larger diameter vessels, finally entering a
broad chamber termed an atrium. From this chamber
water exits the sponge body through a specialized
structure termed an osculum. Such oscula channel water from large portions of a sponge and each sponge is
likely to have several oscula. Water is forced from the
oscula with sufficient force so as to enable materials
102
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
that have passed through the sponge to be transported
far enough away from the sponge that these materials
are unlikely to be recaptured by the flow of water entering that sponge.
Although the description above may suggest that the
structure of the sponge canals was fixed, the system is actually dynamic. The positions of both the incurrent and
excurrent canals, as well as their components, can shift
as the sponge grows or as portions of the feeding canals
are occluded by ingested materials (Mergner, 1966).
Similarly, the mesohyl of freshwater sponges exhibits
rhythmic condensation that assists the choanocyte chamber in pumping water (Weissenfels, 1990).
2. Digestion
A second key feature of sponge feeding involves
the capacity of the cells within the canal network to engulf particles through phagocytosis. Cells ranging from
the porocytes that form the ostia on the sponge surface
to the pinacocytes lining the incurrent canals and the
choanocytes exhibit this capacity for phagocytosis. Digestion and the transport of nutrients within the
sponge body also involve phagocytic activities in which
materials are exchanged between cells. Information on
activities by cells in sponge digestion has been gained
by several detailed studies employing light and electron
microscopy (van Weel, 1949; Schmidt, 1970; Willenz,
1980).
Phagocytosis can occur when a particle makes contact with a surface within the canal system. Food particles large enough to occlude a sponge’s ostia are taken
up directly at the epithelium. Smaller particles are
taken up after contact with the cells lining the canal
walls or with the collars of the choanocytes. Following
initial uptake, food particles are transferred to digestive
cells, termed archaeocytes, which move freely within
the mesohyl. As digestion proceeds, archaeocytes bearing ingested materials move through the sponge interior and transfer nutritional materials to other cells,
such as those involved with reproduction or skeleton
formation.
After the digestion of ingested materials is completed, archaeocytes move to the excurrent portion of
the canal network. Nondigested material is released into
the water flowing out of the sponge by a reverse phagocytic action. Egested particles are then carried by excurrent flow out through oscula at the sponge surface.
Detailed analyses of cell activities in sponge digestion have revealed a surprising degree of complexity.
The time necessary for a particle to traverse the digestive system completely, from initial uptake to release
into an excurrent canal, can vary markedly with the
nature of the food particle and with environmental
conditions. Ordinarily, digestible food particles may be
held within archaeocytes for more than 12 h prior to
release into the sponge excurrent. Under certain circumstances, sponges can sense indigestible particles
and release them after much shorter time periods
(Frost, 1980a). The capacity to speed the processing
time for ingested materials is most evident when a
sponge is faced with a dense suspension of particles. At
such times, food particles may be taken up, cycled, and
released almost immediately (Schmidt, 1970).
The sophistication of the sponge feeding system is
particularly evident in digestive activities because the
rapid cycling of ingested particles can be highly selective. While overabundant particles are being quickly
cycled through the sponge, other food particles, present
in lower concentrations in the feeding suspension, are
held within the sponge for periods that are sufficiently
long to allow normal digestion (Frost, 1980a, 1981).
Sponges, therefore, are selective feeders, not in their
initial uptake of particles as selective feeding is typically defined, but in their ultimate use of these
resources (Frost, 1980b).
A sponge’s capacity to vary particle transit time is
linked to the large number of cells that are operating
within it. Digestive cells are sufficiently numerous so
that, under normal feeding conditions, each cell handles only one ingested particle at a time (Willenz,
1980). Each individual archeocyte then, can be considered as a separately functioning digestive unit with the
capacity to tune its actions to the characteristics of the
food particle that it is processing.
3. Skeleton
The gross structure of freshwater sponges derives
from an interplay between two fundamentally different
components — a mineral skeleton made up of siliceous
structures termed spicules and an organic skeleton
made up of collagen. Collagen binds spicules together
into rigid structures yielding the sponge’s basic framework. All freshwater sponges exhibit skeletal systems
comprised of siliceous spicules and collagen. Marine
sponges exhibit a broader variety of supporting structures (Berquist, 1978).
a. Siliceous Spicules Some major groups of sponges
are unique among multicellular organisms in their use of
silica as a primary component of their skeleton. Freshwater sponge spicules are composed of opalescent silica
that has been laid down along an axial, organic filament
by specialized cells termed sclerocytes. Considerable diversity exists in spicule morphology, and, because much
of this variability is species-specific, their structure plays
a critical role in sponge taxonomy.
Three general classes of spicules are recognized:
(1) megascleres, which make up the main framework of
4. Porifera
103
FIGURE 6
A scanning electron micrograph of megascleres from Spongilla lacustris collected in a lake
with a high concentration of dissolved silica. A spined microsclere is also present. The bar is 10 m in
length.
a sponge; (2) microscleres, which appear to add structural reinforcement for sponge tissues; and (3) gemmoscleres, which form part of the resistant coat of gemmules (see below). Megascleres are needle-shaped
structures (Fig. 6 and Fig. 7) that range in length from
150 to 450 m. Microscleres are similar in form to
megascleres but are usually less than one-fifth their
FIGURE 7
length (Fig. 6). Gemmoscleres are similar in size to microscleres and both exhibit a variety of forms ranging
from needlelike to dumbbell or star-shaped. Any of
these three spicule forms may be smooth or spined
depending on the species. An elaborate terminology has
been developed to describe the varied shapes of spicules.
These terms have been summarized in Boury-Esnault
A scanning electron micrograph of a megasclere of Spongilla lacustris collected in a lake with low
concentration of dissolved silica. The bar is 10 m in length.
104
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
and Rützler (1997), but their use has been kept to a
minimum in this chapter. The fine-scale structure of the
spicules, particularly the gemmoscleres, plays a critical
role in the taxonomy of sponges, and its importance
cannot be overemphasized. The classification of freshwater sponges is based fundamentally on the structure
of gemmule spicules (Penney and Racek, 1868).
b. Collagen The predominant form of collagen in
sponges is spongin, which serves primarily to bind
the spicules of the inorganic skeleton together. A second form of collagen provides smaller-scale structure
within the sponge mesohyl. Finally, collagen, often
combined with gemmoscleres, forms the resistant coat
of gemmules.
4. Reproduction
Both sexual and asexual processes can play major
roles in sponge reproduction. Sexual reproduction involves the activities of numerous, isolated reproductive
cells functioning throughout the body of an active
sponge. Asexual reproduction often involves major
changes in all of the cells within a sponge.
a. Sexual Reproduction Although detailed observations have been made for only a few freshwater species,
it appears that the mode of sexual reproduction is usually gonochoristic, with each separate sponge being entirely male or female. In at least some cases, however,
the gender of a particular sponge is not fixed but appears to depend on environmental conditions. In at
least one species, S. lacustris, sponges have been shown
to switch sex from one year to the next (Gilbert and
Simpson, 1976a). Bisbee (1992) did report one hermaphrodite specimen in a population of S. lacustris in
South Carolina where most sponges exhibited only
male or female elements.
Sexual reproduction is accomplished by specialized
cells that develop within the mesohyl during limited
portions of the year. Reproductive cells are derived from
other, more common sponge cells. Spermatogenesis
occurs within distinct spermatic cysts which appear to
be formed from choanocytes. Egg cells (oocytes) seem
to develop either from archeocytes or choanocytes.
Oocyte growth is dependent upon a variety of nurse
cells that are phagocytized as the egg develops (Gilbert,
1974). Because these events do not occur for extensive
periods, successful sexual reproduction requires synchronous timing of egg and sperm development across
sponge populations in a habitat. This has been documented for some freshwater species (Simpson and
Gilbert, 1973). This requirement leads to spectacularly
synchronized events of extensive sperm production in
some marine species (Reiswig, 1970).
Fertilization occurs when sperm that have been released into the open water by other sponges are
brought into the canal systems of female sponges by
their feeding currents. Sperm are then taken up by the
sponge, probably by choanocytes, and conveyed to an
egg cell where fertilization takes place.
As is the general case within the Porifera, freshwater sponges are viviparous. Larvae undergo extensive
development prior to their release from the mother
sponge. Material for growth is provided by nurse cells.
In the final stages of development, the sponge larva,
termed a parenchymula, contains archeocytes, pinacocytes, sclerocytes, and choanocyte chambers. In addition, its surface is covered by flagellated epithelial cells
which allow the larva to swim upon release.
Once development is completed, larvae are released through the excurrent portion of the feeding
canal system. The larvae swim until they settle onto a
substratum, where they undergo metamorphosis and
quickly develop the structures typical of adult sponges.
b. Asexual Reproduction Asexual reproduction in
sponges may be simple or complex. On one extreme,
clonal development of separate, independently functioning sponges can take place through fragmentation.
The plastic nature of sponge cells makes it possible for
even very small pieces to develop into active, fully functional sponges. Propagation in this fashion occurs commonly in some habitats (Frost et al., 1982).
On the other extreme, most freshwater sponges
typically form gemmules in a complex process of dedifferentiation in which the structures of the active
sponge regress into masses of cells surrounded by resistant coats (Figs. 1 and 8). This process, which often occurs prior to the onset of rigorous environmental conditions, permits sponges to weather periods of stress
and to disperse into other habitats.
The coat of a mature gemmule usually consists of
three collagen layers. In most species, specialized
spicules, gemmoscleres, which are formed only during
gemmule formation, are also embedded within the gemmule coat. The coat of each gemmule is continuous
with a much thinner layer at a single specialized structure, termed a micropyle or foraminal aperture, through
which cells emerge during germination. Within the gemmule are a large number of specialized cells, thesocytes,
which contain numerous yolk inclusions. Thesocytes
can store energy-rich materials which are used for
growth following germination. The aggregation of cells
to form gemmules takes place within the mesohyl.
Gemmules are often present in large numbers within the
vestigial skeletons of previously active sponges.
Completely formed gemmules exhibit low metabolic rates until germination is induced, and are
4. Porifera
105
FIGURE 8
Numerous gemmules (the small whitish spheres) of Spongilla lacustris in a formerly active
fingerlike growth of the sponge.
extremely resistant to environmental stress. Gemmules
of a common freshwater species Ephydatia muelleri
survived exposure to 80°C for more than 9 weeks
(Barbeau et al., 1989) although cold tolerance has been
shown to vary substantially among species (Ungemach
et al., 1997). Gemmules can also survive anoxic conditions for several months (Reiswig and Miller, 1998).
Upon germination, the thesocytes increase their metabolic rate, exit from the gemmule coat through the
micropyle, and differentiate into the varied cells typical
of an active sponge.
At least one freshwater sponge species is also capable of another form of asexual reproduction, budding.
Buds exhibit many of the structures of active sponges,
but in a smaller form that is capable of dispersal within
water (Saller, 1990). They do not, however, exhibit the
same resistance to adverse environmental conditions as
gemmules.
5. Biochemistry
Sponges exhibit a variety of unusual chemical constituents. Some, particularly those identified as secondary metabolites, have been linked with an ability to
deter predation (Pennings et al., 1994). Particularly unusual among chemical features of sponges has been a
major portion of fluorine in the body of a marine
species (Gregson et al., 1979). Most unusual chemicals
have been documented in marine species, but some are
also likely to occur in freshwater species. It has been
suggested that they may provide some defenses against
predation and overgrowth by epizooic organisms for
freshwater sponges (Frost, 1976a).
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
1. Diversity
The diversity of sponges in freshwater is low in
comparison with that in the marine habitats from
which they have evolved. Although the entire phylum
Porifera consists of more than 5000 species (Bergquist,
1978), there are probably fewer than 300 freshwater
species worldwide. In the most recent taxonomic revision of the Spongillidae, the largest family of freshwater sponges, Penney and Racek (1968) list a total of
95 species in 18 genera . While it is certain that there
are additional, undescribed species, particularly in
tropical regions that have not been thoroughly investigated (e.g., the Amazon Basin; Volkmer-Ribeiro, 1981),
it is clear that freshwater forms represent a small group
within the sponge phylum.
In turn, North American freshwater sponges
represent only a restricted subset of the world’s freshwater species. Overall, within both the Spongillidae
106
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
and Metaniidae, we have generated a list of 27 clearly
distinct species in 11 genera that appear to be valid
reports for the portion of North America north of
Mexico and the Caribbean.
2. Distribution
From the broadest perspective, the distribution of
sponge species is influenced by biogeographic factors.
On an intermediate level, the general environmental
conditions that exist within a lake or stream will control the presence of a particular sponge species there.
Finally, conditions such as light, substrata, and wave
action will determine sponge distribution within a particular water body.
The freshwater sponge species in North America
exhibit a variety of distribution patterns (Penney, 1960;
Penney and Racek, 1968). Several species have been
reported throughout the United States and Canada,
while others have been observed in only one or a few
habitats. In some cases, a broad distribution in North
America represents only a subset of a more extensive
range. Ephydatia fluviatilis and Eunapius fragilis
are distributed worldwide, while Spongilla lacustris,
Ephydatia muelleri, and Trochospongilla horrida occur
throughout temperate regions of the Northern Hemisphere. Some species exhibit more restricted crosscontinental distributions. Anheteromeyenia ryderi occurs primarily in eastern North America but has also
been reported once in Central America and from several locations in Western Europe (Økland and Økland,
1989). Other species are broadly distributed throughout North America, but have not been observed
on other continents (e.g., Duosclera mackayi and Trochospongilla pennsylvanica). Of the remaining North
American species, most appear to be restricted to limited regions within Canada and the United States; and,
in the most extreme cases, several species have been reported from single locations.
It is important to note that this chapter provides
limited coverage of North America. It considers only
sponges in Canada and the United States. Limited information on Mexican sponges is reported in Rioja
(1953). Smith (1994) presented the first report of a
freshwater sponge from the West Indies. Abundant
sponges have been observed in a wetland in Costa Rica
(T. Frost, personal observation), and it is certain that
there are sponges throughout Mexico and Central
America.
Within Canada and the United States, there have
been numerous reports on the occurrence of sponges in
provinces and states or regions within them. Some recent examples include reports for Alberta (Clifford,
1991), Arizona (Sowka, 1999), Connecticut (De Santo
and Fell, 1996), eastern Canada (Ricciardi and Reiswig,
1993), western Montana (Barton and Addis, 1997), and
southern Lake Michigan (Lauer and Spacie, 1996). Penney (1960) presented an extensive review of earlier reports on freshwater sponges. Despite these reports, information available on the occurrence of sponges does
not match their overall distribution.
The apparently limited ranges of some species may
result more from a lack of observations rather than actual restrictions in sponge distributions. Accepting this
limitation, however, there appear to be some general
trends in the distribution of sponges across North
America. For example, relatively few species of
sponges have been reported in western regions, and
there appears to be a general east-to-west decrease in
sponge diversity. A similar trend occurs between
northern and southern regions, with more species reported from the northern United States and southern
Canada than from the more southern regions of the
continent. In some situations, the limited distribution
of a particular species may involve adaptations to climatic conditions. In many other cases, biogeographic
factors, particularly limitations on dispersal, are likely
to control regional distribution patterns. There are
some notable exceptions to limitations on dispersal,
however (e.g., Jones and Rützler, 1975, Økland and
Økland, 1989).
Within regional distribution patterns, the factors
that regulate the occurrence of a species in a particular
lake or stream are less well understood. While
Spongilla alba appears to be restricted to brackish
water habitats (Poirrier, 1976), other species are influenced by a variety of environmental factors. Jewell
(1935, 1939) provided the most detailed quantitative
information available on this topic. In comparing
species across lakes in northern Wisconsin, she found
considerable variation in each species’ habitat requirements. Some species were restricted to a narrow subset
of environmental conditions, while others occurred
throughout a broad range of habitats. Chemical factors
were correlated with the distributions of some species;
but for other species, more recent analyses indicate that
there is wide tolerance of chemical conditions (Colby
et al., 1999). In addition, some species were favored by
flowing water, while others were more typically associated with standing water. In general, it is difficult to
pinpoint any specific factors that control the distribution of sponges. Many species are sufficiently broad in
their distribution so that many lakes can be expected
to contain at least some small specimens. This is particularly true in some regions, e.g., northern Wisconsin,
where sponges occur in nearly every lake. In some
cases, particularly in smaller lakes, large populations
of several sponge species may be common. Aside
from cases of geographic isolation, sponges would be
4. Porifera
excluded only from habitats with frequent physical disturbances, with high pollution levels, or with high
loadings of silt or particulates that can clog their feeding systems (Harrison, 1974). Not all pollutants affect
sponges, however. They appear to be tolerant of high
levels of some contaminants, particularly heavy metals
(Richelle et al., 1995).
Within a lake or stream that is suitable for growth,
sponge distribution is controlled by finer scale environmental features. For most sponges, a hard substratum
is essential for growth, and the absence of a surface on
which to grow will limit the distribution of sponges
even where other conditions are highly favorable. Only
a few species, notably Spongilla lacustris and, to a
lesser extent, Ephydatia muelleri, can grow in soft sediments. In rivers and streams, sponges are excluded
from regions with high flow due to physical disruption.
In lakes, sponges can be limited in shallow waters by
wave action or ice scour (Bader, 1984). In deeper waters, sponges can be limited by low oxygen or by colder
temperatures. Some species with symbiotic algae may
be limited by light availability, particularly where water
is darkly stained by dissolved organic materials, but
species with less dependence on symbionts may thrive
in such habitats.
B. Reproduction and Life History
The annual life-history pattern for most freshwater
sponges involves periods of active growth alternating
with dormancy and includes both asexual and sexual
reproductive processes. Transitions to, and from, dormant stages usually involve the asexual processes of
gemmule formation and hatching. Asexual reproduction also occurs during active growth periods, both in
the formation of gemmules and in the generation of
separately functioning sponges through fragmentation.
Sexual reproduction occurs only for a limited time
during periods of active growth. Key features of
the freshwater sponge life cycle are summarized in
Figure 9 and discussed in detail below. Overall, reproduction in sponges involves responses to two distinct
problems, propagation within a single habitat and dispersal among habitats. Within-habitat processes are
discussed separately below prior to a consideration of
dispersal.
107
either situation, sponge feeding ceases and respiratory
activities are greatly reduced. Most studies of dormancy in sponges have focused on transitions involving
gemmules.
Despite the common role of gemmules in dormancy, their presence does not in itself necessarily signify a transition to a dormant life-cycle period. In many
cases, particularly where sponges are large, a few gemmules are produced during most of the active season.
Most freshwater sponges undergo a period of dormancy at some time during the year, typically during
periods of environmental stress. In temperate habitats,
dormant periods occur most commonly during winter.
This is the case for the majority of sponges in North
America. However, at lower latitudes this pattern can
be reversed with dormancy occurring during hot,
summer periods and active growth taking place during
cooler seasons (Poirrier, 1974; Ilan et al., 1996).
Dormant periods have also been observed in response
to drying in ephemeral habitats. While dormancy is a
common feature of freshwater sponge life cycles, it
does not always occur; some sponges maintain active
growth even under winter ice cover (Frost, personal
observation).
The timing of dormancy varies both among species
within particular habitats and among habitats for
individual species. In Lake Pontchartrain, Louisiana,
Spongilla alba exhibits a typical pattern of gemmule
formation during winter months (Poirrier, 1976),
while a co-occurring population of Ephydatia fluviatilis
exhibits a reciprocal pattern of summer dormancy
(Harsha et al., 1983). At the same time, populations
1. Dormancy
In most cases, dormant periods for freshwater
sponges are characterized by the complete transformation of all active tissue in a sponge into gemmules
(Simpson and Fell, 1974; Simpson, 1984). More rarely,
tissue regression yields a growth form that functions
as a gemmule without its specialized structures. In
FIGURE 9
General features of the annual life history of freshwater
sponges (see text for detailed explanations).
108
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
of E. fluviatilis in more northern habitats form dormant stages in winter. S. lacustris populations in New
Hampshire gemmulated during winter and were active
during summer (Simpson and Gilbert, 1973), while
population in South Carolina were inactive during
summer periods (Bisbee, 1992). Sexual reproduction
occurred at different times during the seasonal cycle
in these areas as well. In Little Rock Lake, Wisconsin,
S. lacustris undergoes a winter dormancy period while
co-occurring populations of Corvomeyenia everetti and
Ephydatia muelleri maintain active growth during
the entire winter. Populations of E. muelleri in nearby
Wisconsin lakes undergo typical winter dormancy.
The transition from an active to a dormant state for
sponges usually appears to be cued by environmental
factors, such as changing temperatures or declining water levels. In some species this transition may also involve a complex response to previous growth conditions
(Gilbert, 1975). Within a region, the timing of gemmulation is nearly synchronous across habitats for all populations of a species that exhibit dormancy (Frost, personal
observation). In contrast, different species within a region can vary substantially in the seasonal timing of dormancy and in the rate at which the transition from active tissue to gemmules proceeds. In Mud Pond, New
Hampshire, Trochospongilla pennsylvanica begins forming gemmules in mid-August when water temperatures
are at their maximum while S. lacustris does not form
gemmules until October when water temperatures reach
10°C (Simpson and Gilbert, 1973).
Sponge species also vary in the factors that induce
their release from dormancy (Simpson and Fell, 1974).
Temperature appears to be the primary environmental
cue that induces gemmule hatching in temperate habitats. Gemmules of some sponges hatch while water
temperatures are at near-winter levels suggesting that
another cue such as photoperiod or the availability of
light is operating.
In some species, gemmules exhibit a form of diapause in which a certain period at cold temperatures is
necessary before hatching can occur (Simpson and Fell,
1974). Such a delay prevents hatching during short,
warming periods prior to the onset of winter. Hatching
at such times could lead to major population reductions if time or energy was insufficient for a second formation of gemmules. Not all species form diapausing
gemmules and, in these cases, hatching will take place
during any period of increasing water temperature. The
occurrence of diapause appears linked to the timing of
gemmule formation. Species that form gemmules later
in the year are less likely to exhibit diapause. While
there are obvious costs to hatching at the wrong
time, there may also be advantages to being able to
hatch quickly without diapause under some circum-
stances. Studies of dormancy in freshwater sponges
have generally focused on cold-water populations, and
less is known about the factors that control the transitions to, and from, dormant stages when they occur in
response to hot or dry conditions.
In general, while it is clear that the formation of dormant stages plays a crucial role in the overall life history
of most sponges, substantial variability is apparent in:
(1) the morphological forms involved in dormant stages;
(2) the timing of transitions to, and from, dormant periods; and (3) the occurrence of diapause. This variability
suggests that the overall value of dormancy and the successful strategies by which it can be employed differ
markedly among habitats and that sponge responses to
such differences involve a diversity of adaptations.
2. Growth, Reproduction, and Dispersal
within Habitats
Nondormant periods in the freshwater sponge life
cycle appear to be characterized by continuous growth.
Although quantitative studies have been limited, sponge
growth in some habitats can be extremely prolific. In a
pond in New Hampshire, 10 mg dry mass of gemmule
tissue of S. lacustris that hatched in spring produced an
average of 18 g of active sponge tissue just prior to
gemmule formation the following fall (Frost et al.,
1982). Ephydatia fluviatilis exhibited comparable prolific growth rates in a European stream (Pronzato and
Manconi, 1995). Growth is largely indeterminate for
individual sponges, which may reach surprisingly large
sizes. In some optimal habitats, for example, a single
specimen of S. lacustris, consisting of intertwined fingers (as in Fig. 1), can occupy more than a cubic meter
of space (Frost, personal observation).
Sexual reproduction for most freshwater sponge
populations occurs synchronously throughout a habitat
in a 1 – 3 month period following hatching from dormant conditions (Simpson and Gilbert, 1973). Synchrony across a population is particularly important,
because otherwise the sperm released by male sponges
would have a low probability of successfully fertilizing
an egg (Gilbert, 1974).
The motile larvae produced by sexual reproduction
in freshwater sponges can play an important role in the
establishment of new sponges. In particular, the colonization of new substrata will often depend on such
motile forms. While settling on an appropriate substratum is clearly critical for the successful growth of a
new sponge, little information is available on substrate
choice by settling sponge larvae. Studies of a variety of
marine organisms suggest, however, that substrate selection by freshwater sponge larvae may be possible.
Fragmentation of intact specimens into separately
functioning units may also play an important role in
4. Porifera
freshwater sponge life cycles. In some cases, such fragmentation occurs during periods of tissue regression
and may not lead to growth on new substrata. In other
habitats, however, particularly those with large
amounts of aquatic vegetation to which sponges can attach, fragmentation and growth may occur repeatedly
during an active season leading to dispersal throughout
a habitat (Frost et al., 1982).
Although gemmules are frequently associated with
dormant periods, they may also be present in freshwater species during periods of growth. Many sponge
species routinely contain a few gemmules within otherwise active tissue, particularly in areas of thick growth.
In an extreme case, S. lacustris develops specialized,
summer gemmules which have much thicker coats than
the gemmules formed for overwinter periods (Gilbert
and Simpson, 1976b). These investigators suggested
that summer gemmules may serve in a “bet-hedging”
strategy against some environmental catastrophe which
could occur prior to more extensive gemmule formation. It seems likely that summer gemmules may also
serve in dispersal among lakes or streams.
3. Dispersal Among Habitats
As is the case for most freshwater organisms, an
ability to disperse among different habitats is critical
for the continued success of sponges in freshwater environments. Most lakes and streams are ephemeral from
an evolutionary perspective. Nearly all lakes in North
America are less than 20,000 years old (Frey, 1963), a
brief period compared to the 100 million year history
of freshwater sponges (Racek and Harrison, 1975).
While there has been little documentation of dispersal by freshwater sponges among habitats, it seems
likely that gemmules are the primary agent for such
movement. The two other life-cycle stages that might
function in dispersal among habitats, larvae and
sponge fragments, possess neither the structural nor the
physiological mechanisms necessary for movement out
of water or across significant distances. It is conceivable that these forms, particularly the larvae, could
function in short-range dispersal among connected water bodies, but their fragile structure would preclude
any out-of-water transport. Gemmules, on the other
hand, with their resistant coats and ability to withstand
harsh environmental conditions appear to be well
suited to dispersal among habitats. Ricciardi and
Reiswig (1993) have reported rafting of S. lacustris
gemmules during flooding in the Ottawa River.
C. Ecological Interactions
As is the case with all organisms, the size of
a freshwater sponge population, and its subsequent
109
impact on an ecosystem, results from an interplay of
growth and loss processes. Growth is regulated by the
availability of nutrients and suitable habitat. Losses are
influenced by physical environmental conditions and,
potentially, by interactions with other organisms.
Although quantitative studies are few, it appears that
the dynamics of freshwater sponge populations are
mediated largely by the availability of suitable microhabitats within a lake or stream. Where substrates for
sponge growth are short lived or where physical disturbances are frequent, growth and loss rates can be
extremely rapid and may operate primarily in a density
independent fashion. Under such circumstances, sponge
populations would be regulated by an interplay
between the physical environment and their own
growth and life history processes. Concomitantly, interactions with other organisms would exert only minimal
influences on sponge population processes. In contrast,
where substrates are permanent, sponge populations
appear more likely to be regulated by intra- or interspecific biotic interactions (Jackson and Buss, 1975). In
many sponge habitats where growth on permanent
substrates is rare, such biotic interactions could be
expected to exert only a secondary role in their population dynamics.
1. Availability of Substrates
The development of freshwater sponge populations
usually depends on the availability of hard substrates.
Most species must be attached to a substrate in order
to grow. In addition, all species seem to depend on the
presence of some solid surface for the successful
settling and growth of their free-swimming larvae. A
variety of surfaces are suitable. Examples range from
boulders or exposed bedrock to the branches of fallen
trees to the leaves and stems of aquatic macrophytes.
Human built structures in water, such as bridge foundations, dams, spillways, and floats, can also serve as
suitable surfaces for sponge growth. Potential substrates vary markedly in their permanence relative to
the longevity of sponges, and these differences, in turn,
have important consequences for sponge population
dynamics.
Among North American species, only S. lacustris
has been routinely observed growing out of soft sediments. A quantitative investigation of its population
dynamics (Frost et al., 1982), however, illustrated a
crucial role for substrates even where sponges did not
depend upon such substrates for their growth. In Mud
Pond, New Hampshire, S. lacustris grows either directly from soft bottom sediments or attaches to any of
several species of aquatic macrophytes which exhibit
large summer populations, but which die back completely before winter. Due to the near absence of any
110
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
permanent substrates in Mud Pond, gemmulated
sponges there overwinter almost exclusively in sediments on the pond bottom. Successful hatching in
spring depends upon the presence of enough gemmulated tissue to grow out of these sediments, and many
sponges are not large enough to make this transition.
Most sponges are unsuccessful in hatching, and spring
populations are sparse. Hatching from sediments,
therefore, is the limiting step in the Mud Pond population’s dynamics. Despite remarkable growth during
summer (a 1,000-fold increase in biomass), the population of S. lacustris in Mud Pond was maintained at
nearly constant levels throughout five years of observations. Essentially, the population is maintained in a
density-independent fashion by a combination of the
lack of permanent substrates and the difficulty of gemmule hatching from soft sediments.
Sponge population dynamics appear to be substantially different in habitats where more permanent
substrates are available. Populations of Ephydatia
fluviatilis growing on hard substrates in a river on
Sardinia, Italy exhibited growth rates that were still
high, but were substantially lower than those reported
for S. lacustris (Pronzato and Manconi, 1995). Populations were persistent on rocks over several years making transitions from active to gemmulated tissue. They
were substantially disrupted, however, by a major flood
but did recolonize habitats fairly quickly (Pronzato and
Manconi, 1995). In general, there appear to be periods
of substantial population growth with substantial
degeneration during other parts of the year, a
process that can be categorized as density-independent
(Pronzato and Manconi, 1995). Finally, in Mary Lake,
Wisconsin, S. lacustris grows attached to trees that
have fallen into the lake and sponges repeatedly make
the transition from active tissue to gemmules and back
to active tissue within the same skeleton over the
course of several years. In contrast with the large fluctuations in sponge density observed in Mud Pond, it
seems that the populations of S. lacustris in Mary Lake
exhibit higher densities throughout the year and lower
growth rates during the course of the summer than
those seen in Mud Pond (Frost, personal observation).
Under such circumstances, population dynamics can be
characterized primarily as density-dependent.
Overall, the availability of substrates exerts a
major control over sponge population dynamics. For
many species, the lack of substrates limits growth
completely. But, even where species are not wholly
dependent on substrates, their nature and availability
can exert a major influence over sponge population
dynamics. The longevity of an individual sponge, including repeated transitions between active and gemmulated states, would appear to be directly related to
the permanence of its substrate. In addition, it seems
likely that the importance of both intra- and interspecific biotic interactions in sponge population dynamics
will be influenced by the time that a particular substrate has been available for colonization. Only on
long-lived substrates would sponges be likely to reach
sufficient densities to interact with each other.
2. Nutritional Ecology
When suitable habitats are available, sponges are
capable of substantial growth. In some cases, these high
growth rates appear to be linked primarily to a sponge’s
efficient particle-gathering mechanisms. In other situations, however, sponges have an alternative means of
obtaining resources. Many freshwater sponges contain
large numbers of intracellular algae. They form symbiotic associations that combine autotrophy and heterotropy. In addition, because sponges require substantial amounts of silica for their spicules, population
growth and maintenance have the potential to be influenced by silica availability.
a. Sponge Feeding Although there have been several detailed studies of the mechanisms by which
sponges feed and of the resources they consume (Frost,
1987), much less is known about the influence of food
availability on sponge growth under natural conditions. The rates at which sponges filter water can be extremely high. During summer, specimens of S. lacustris
typically filtered more than 6 ml/h per miligram dry
mass of tissue (Frost, 1980b). At this rate, a finger-sized
sponge filters more than 125 L per day . Because a
sponge’s feeding effectively removes all particles ranging in size from bacteria to large algae from the water
it filters, these rates suggest that sponges may exhaust
available food under some circumstances. Food limitation would be particularly likely in still water or where
particulate resources are sparse. Pile et al. (1997)
demonstrated that such depletion occurred in Lake
Baikal, Russia, where sponges consumed a variety of
picoplankton (cells 2 m in diameter) in substantial
quantities.
b. Algal Symbionts In many cases, freshwater
sponges are bright green due to the presence of large
quantities of chlorophyll contained within extensive
populations of algal symbionts (Gilbert and Allen,
1973a, b). This situation is so common that sponges
are frequently mistaken for plants in freshwater
habitats. The algal symbiosis in freshwater sponges is
representative of a large number of algal – invertebrate
associations that are common in marine and freshwater habitats (Reisser, 1984). In these mutualistic
4. Porifera
associations, the nutritional processes of algae and invertebrates are closely coupled. In freshwater sponges,
and in many other invertebrates, algae are maintained
endosymbiotically within the cells of their host.
Algal – invertebrate symbioses combine autotrophic
processes with normal animal heterotrophy. In the resulting, mixotrophic nutrition, symbiotic algae provide
photosynthetically fixed carbon to the invertebrate host
which, in turn, supplies nutrients such as nitrogen or
phosphorus or carbon dioxide to the algae. This combination has proved particularly successful in nutrient
poor conditions, such as coral reefs (Muscatine and
Porter, 1977) but occurs over a broad range of habitats.
Algal symbionts play a major role in the growth of
some freshwater sponges. Contributions from symbiotic algae accounted for 50 – 80% of the growth of S.
lacustris in Mud Pond, New Hampshire (Frost and
Williamson, 1980). The contribution of algae was determined by maintaining sponges in situ under darkened conditions, which led to the loss of their algae.
Comparisons were then made between the growth of
such aposymbiotic sponges and that of normal green
sponges maintained under lighted but otherwise identical conditions. In similar experiments in Little Rock
Lake, Wisconsin, S. lacustris and two other sponge
species were incapable of any growth and died after a
few weeks under darkened conditions (Frost, personal
observation). Sand-Jensen and Pedersen (1994) also
demonstrated a substantial contribution by symbiotic
algae to the growth of S. lacustris in a Danish stream.
Not all sponges depend on algal symbionts, however.
Some species can live in darkness, where their nutrition
is based solely on heterotrophic processes.
The factors that control the proportional contribution of autotrophic and heterotrophic processes to
overall sponge nutrition are less clear. That sponges
failed to grow under darkened conditions in only some
habitats indicates that the relationship between a
sponge and its algal symbionts can range from facultative to obligate depending on environmental conditions. Even for species that sometimes depend to a
large extent upon their algae, the amount of algal
chlorophyll within a sponge (a rough measure of the
potential contribution of its algal symbionts) varies
markedly, both among habitats for a species and
among species within a single habitat. In a survey of
northern Wisconsin lakes, the average concentration of
algal chlorophyll in S. lacustris varied among lakes by
more than a factor of three (Frost, personal observation). Similarly, in Little Rock Lake, Wisconsin, three
sponges exhibited consistent species-specific differences
in their chlorophyll content that were coupled with significantly different rates of both photosynthesis and
feeding (Frost and Elias, 1990). These differences
111
occurred even though the three sponge species shared
identical environmental conditions.
The symbiotic relationship between sponges and algae differs fundamentally among sponge species. Some
species depend, to a large extent, on algae for growth,
while others exhibit little dependence. These differences
among species may involve specializations to particular
environmental conditions, in that sponges that contain
large amounts of algal chlorophyll may be capable of
growth only where light is readily available. Comparing
habitats, it seems likely that sponges would vary the
density of symbionts that they contain in response to the
relative availability of light and particulate resources.
However, this relationship is not simple. Detailed examinations of several sponge species indicate that increasing
particulate resources may favor higher rather than lower
concentrations of algae (Frost and Elias, 1990).
Until recently, it had been thought that all symbiotic associations of algae and invertebrates in freshwater involved the presence of one group of green alga
termed zoochlorellae (Reisser, 1984). Invertebrates
hosting this alga include protozoans, hydra, flatworms,
clams, and sponges. It has now been determined, however, that at least one freshwater sponge species,
Corvomeyenia everetti, and likely its congener C.
carolinensis, contain a yellow-green alga as their symbiont (Frost et al., 1997). Volkmer-Ribeiro (1986) has
suggested that the Corvomeyenia species have an evolutionary history that is long distinct from sponges
with green-algal symbionts. These distinct histories
combined with numerous overall similarities between
sponges with green and yellow-green symbionts indicate that two, closely convergent symbiotic associations
have evolved separately in the freshwater sponges.
Morphological affinities between marine algae and the
yellow-green symbiont suggest that the symbiotic association in Corvomeyenia may have developed in an
ancestral marine form and been maintained during a
subsequent invasion of freshwater. It is important to
note that differences between zoochlorellae and yellowgreen algae cannot explain the observed differences in
algal contributions to overall sponge nutrition described above. Substantial differences in the contribution of algae have been observed in comparisons of
sponge species that contain only green algal symbionts.
The variety of ecological interactions between
sponges and algae and the complexity of their evolutionary history indicate that algal symbioses have been
important in the diversity and distribution of freshwater sponges.
c. Potential Alternative Nutritional Modes Additional means of gathering food resources have been
documented recently for some marine sponges. Vacelet
112
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
and Boury-Esnault (1995) have described a deep-sea
sponge that lacks a typical sponge water-processing
system and feeds on small animals with a mechanism
that can be considered as carnivorous. However, some
carnivorous sponges near deep-sea hydrothermal vents
exhibit a symbiotic relationship with methanotrophic
bacteria (Vacelet et al., 1995). Neither nutritional
mode has been documented in freshwater species,
but their existence in marine habitat raises interesting
possibilities.
d. Influence of Silica on Skeleton Structure The
availability of silica within a habitat may exert both
direct and indirect effects on sponge growth. As discussed previously, some sponges are restricted to high
or low silica habitats. However, some species are distributed across habitats exhibiting a broad range of
silica conditions, where they exhibit major responses
to its availability. For example, in populations of S.
lacustris in different lakes in northern Wisconsin, the
total amount of biogenic silica and both the number
and morphology of megascleres vary directly with silica
concentrations in the lake (Frost et al., in preparation).
Total biogenic silica and spicule width (Fig. 6) decrease
as less silica is available. In contrast, however, the number of megascleres increases as their width decreases
(Fig. 7). With this combination of changes, the total
surface area of megascleres remains constant as silica
decreases — a situation that increases the structural
support gained per unit of available silica. This suggests a direct and complex response by sponges to different availabilities of silica.
Responses to low silica conditions are related to
changes in the stiffness and growth form of sponges.
Specimens of S. lacustris from high silica habitats are
much stronger and more resistant to breakage than
those growing where silica is low. Sponges in low silica
habitats, therefore, may be limited to growing closely
attached to substrates or in waters with little wave
action. Also, because spicules may play a role in the resistance of sponges to predation (see below), populations in low silica habitats may be more vulnerable to
consumption.
3. Biotic Interactions
a. Competition for Substrates The availability of
substrate plays a crucial role in the population dynamics of freshwater sponges. As such, competition for
surfaces would seem to have a potentially important effect on sponge distribution and abundance. A sponge
has the potential to interact with a substrate any time
that its growth leads to contact with another sponge
or any other organisms. There are reports of significant
interactions for a substrate within, and between,
sponge species, as well as between sponges and other
organisms. Such interactions do not appear to occur
commonly, however, and the role of competition for
a substrate in the regulation of freshwater sponge
populations remains largely unknown. In part, this is
due to the minimal attention that interactions for substrates have received in freshwater in general. In addition, because freshwater sponge populations are often
limited by density-independent processes, the abundance of sponges within a habitat is usually sparse with
little potential for interactions on substrates.
When specimens of same sponge species grow into
contact with each other on a substrate, two distinctly
different results have been observed (Van de Vyver,
1970). In some cases, the separate sponges will form a
single, functionally integrated unit. In other instances, a
distinct structural barrier is erected in areas of contact
and the two sponges continue to function as completely
separate units. The genetic relationship between
sponges appears to control whether they will merge or
develop a barrier. Separate sponges grown from fragments or gemmules taken from the same original
sponge will always merge with each other. In contrast,
experimental manipulations have indicated the existence of distinct strains within a sponge species which
will not merge with each other (Van de Vyver and
Willenz, 1975). Studies of marine sponges originally
suggested that only sponges that are genetically identical would merge with each other (Niegel and Avise,
1983). However, subsequent analyses have indicated
that, while the probability of merging appears to be a
function of the genetic similarity of sponges, specimens
need not be genetically identical in order to join into a
single unit (Stoddart et al., 1985). Such fusion interactions in sponges operate at a cellular level (Van de
Vyver, 1979) and have attracted the attention of researchers interested in the basic process of cell – cell
recognition. These studies have direct implications for
the understanding of immune-response systems in more
complex invertebrates and in vertebrates (Smith and
Hildemann, 1984).
Interactions for substrates among different sponge
species have been thoroughly documented in marine
systems while they have received little attention in
freshwater. Marine studies have demonstrated a complex hierarchy in which certain sponge species can
completely overgrow and kill other species (Jackson
and Buss, 1975). These interactions have a major
influence on overall community ecology on substrates
(Jackson, 1977). Observations of interspecies interactions in freshwater sponges have been limited to
instances where sponges have been shown to develop a
nonmerging front similar to that described above for
intra-species interactions (Fig. 1). In habitats where
4. Porifera
substrates are extremely long-lived, it seems likely
that some sponge species may be able to completely
overgrow other sponges, but the significance of such
interactions in freshwater habitats remains to be
demonstrated.
Interactions for substrates between sponges and
other organisms had likewise received little attention in
freshwater. Recently, however, sponges have been
shown to overgrow and kill the zebra mussel, Dreissena
polymorpha, a species whose recent invasion of North
America via the Laurentian Great Lakes has been extensive and has raised substantial concerns (Ricciardi et al.,
1995). Unfortunately, sponge overgrowth does not appear to be a solution to zebra-mussel related problems.
More generally, for attached organisms overgrowth by
sponges must be a factor in other freshwater habitats.
The spreading of a sponge across any surface must involve the overgrowth of microscopic algae and bacteria.
Sponges often encrust on aquatic macrophytes, but do
not seem to harm the plants. In a few cases, freshwater
sponges have been observed to overgrow and kill
colonies of bryozoans (Frost, personal observation).
The overall importance of sponge overgrowth for other
organisms remains to be assessed.
b. Predation and Infaunal Organisms Conspicuous
predation upon sponges, in which major portions are
consumed by a larger organism, is rare. A variety of
smaller organisms consume portions of sponges, but
they appear to act as grazers or parasites, leaving the
sponge that they consume largely intact. Such species
that feed on sponges represent a subset of the diverse
infaunal community that typically occurs in freshwater
sponges. Other infaunal organisms appear only to
make use of the structure provided by a sponge in a
commensal relationship.
Resistance to predation by larger organisms appears to be characteristic of the Porifera in general
(Randall and Hartman 1968; Resh, 1976). Mechanical
and chemical defenses combine to make sponges resistant to predation. Spicules provide mechanical deterrence to predators; these sharp spines are thought to
act as an irritant to its mouth and digestive system. The
effectiveness of spicules in deterring predation was evident in experimental tests in which snails would not
consume S. lacustris from habitats where it contained
robust spicules, but would consume specimens from
habitats in which low silica availability limited the size
of spicules (T. Frost and A. Covich, personal observation). Chemical deterrence to predation has been attributed to a variety of toxic and pharmacologically active
compounds that are well documented in marine
sponges (Cecil et al. 1976) and are also likely to occur
in freshwater species.
113
Despite the general effectiveness of sponge defense
mechanisms, a few organisms prey effectively upon
sponges. In some cases sponge predators have developed a nearly complete dependence on sponges. The
few large predators on freshwater sponges include
fishes, crayfishes, ring-necked ducks and, possibly,
snails. Extensive predation by fish on sponges has been
documented in Africa (Dominey, 1987) and South
America (Volkmer-Ribiero and Grosser, 1981), but is
not common in North America. Crayfish were implicated as causing major reductions in a sponge population in a Massachusetts stream (Williamson, 1979), but
widespread consumption of sponges by crayfish has
not been reported. Juvenile ring-necked ducks have
been reported to sometimes depend on sponges as a
food source (McAuley and Longcore, 1988). Snails will
consume and can subsist on freshwater sponges under
laboratory conditions (T. Frost and A. Covich, personal observation); but here too, their significance as
predators under natural conditions remains unknown.
Numerous small invertebrates reside within, or attached to, freshwater sponges. Examples include protozoans, oligochaetes, nematodes, rotifers, bivalves, water mites, and aquatic insects (Berzins, 1950; Resh,
1976; Volkmer-Ribiero and De Rosa-Barbosa, 1979).
A variety of relationships exist between these infaunal
organisms and their sponge hosts. Some infaunal
species feed upon sponges while others use the sponge
body as a habitat. In some cases the relationships are
obligate, at least for certain life cycle stages, and such
species may show a striking degree of specialization to
the sponge host. Other organisms are only occasionally
associated with sponges.
Several groups of aquatic insects typify organisms
with obligate, specialized feeding on freshwater
sponges. Numerous species within three insect orders,
the Diptera, Neuroptera, and Trichoptera, depend on
freshwater sponges during major portions of their life
cycle (Resh, 1976). One neuropteran family, the Sisyridae, is so commonly associated with sponges that its
members are referred to as Spongilla Flies. Not all relationships between insects and sponges are obligate.
Some insect genera that contain obligately, spongefeeding species also contains species that are only occasionally or never associated with sponges (Resh et al.,
1976). The variety of interactions evident between
sponges and related insect species indicates that the
evolutionary history of insect – sponge relationships is
diverse and complex.
Obligate relationships with freshwater sponges
have also been documented for water mites (Proctor
and Pritchard, 1990) that apparently depend on the
sponge primarily for structure. The water mites live
and lay their eggs within the feeding canal system of
114
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
the sponge. These and other organisms that live within
sponges (e.g., clams, insects and protozoans) may be
taking advantage of the lack of predation on sponges.
Spicules and chemical defenses may provide a general
refuge against predators on infauna as well as sponges.
The community ecology of the infaunal organisms
themselves is also complex. The presence and abundance of a particular infaunal species results from an
interplay of its own life cycle, the life cycle of the host
sponge and potentially, the abundance of other infaunal organisms. Data from northern Wisconsin sponge
populations indicate that most other infaunal species
are rare or absent during periods when water mites are
abundant within sponges (J. Elias and T. Frost, personal observation). Again, this is an area that has received little attention.
tative of sponges in a variety of other lakes and
streams. Sponge populations in some northern Wisconsin lakes, for example, appear to be substantially larger
than those in Mud Pond (T. Frost, personal observation). At the same time, sponges in many ecosystems
represent only a minor component when compared
with other organisms.
Even when sponges account for substantial portions of an ecosystem’s primary and secondary production, they may not interact directly with higher trophic
levels. The lack of predation on sponges can lead to an
accumulation of material in sponge biomass that is not
passed directly up the food chain. Interaction of sponge
with other ecosystem components may be indirect, operating primarily to limit the availability of materials to
other organisms within the food web.
4. Functional Role in Ecosystems
5. Paleolimnology
In most lakes and streams, populations of freshwater sponges are sufficiently sparse that they are unlikely
to play a major role in ecosystem function. However,
there are marked exceptions to this general pattern. In
some habitats, sponges are the dominant component of
the benthic community and exert a substantial influence on total nutrient cycling and primary production.
Quantitative analyses of freshwater sponge populations
are rare, but a detailed study of one population illustrates the potentially major impact of sponges on
ecosystems (Frost, 1978; Frost et al., 1982). At the
peak of its population size in early October, S. lacustris
in Mud Pond, New Hampshire reached an average biomass of greater than 3.5 g dry mass per square meter.
At this density, the total sponge population in the pond
would filter more than ten million liters of pond water
per day, a rate that would cycle a volume equivalent to
that of the entire pond every 7 days. Since the sponges
are capable of removing most phytoplankton and bacteria from the water that they filter, their potential impact on Mud Pond is substantial. Because of their algal
symbionts, sponges can also account for a substantial
portion of the primary production in Mud Pond. During late summer and fall, the chlorophyll within the
sponges is approximately equivalent to that in the entire phytoplankton community (Frost, 1978; Frost and
Williamson, 1980).
Sponges also play a major role in flowing-water
ecosystems. Mann et al. (1972), in a detailed study of
the River Thames in England, found that sponges accounted for nearly 40% of the total production by benthic animals.
Clearly, sponges can be a major component of the
biota of some ecosystems. While detailed quantitative
studies are rare, observations indicate that the populations in Mud Pond and the River Thames are represen-
The siliceous nature of sponge spicules makes them
resistant to decomposition under most circumstances.
As such they are usually well preserved in lake sediments and have the potential to serve as a useful tool in
indicating historic lake conditions. In situations where
specific habitat requirements can be defined for a
species (Jewell, 1935), the presence of its spicules in the
sediments of a lake from which it is currently absent
can indicate changes over time in the environmental
conditions of that lake. Likewise, even when a sponge
species is present throughout an extended period
within a lake, quantitative changes in spicule morphology over time can indicate shifts in the availability of
silica within that lake. For example, lakes throughout
northern Wisconsin appear to have exhibited a more
than 50% reduction in silica availability over the last
12,000 years (Kratz et al., 1991). Paleolimnological investigations employing sponge spicules are a potentially
fruitful area for study (Harrison, 1986). Studies in 28
Connecticut water bodies revealed that most lakes had
maintained the same sponge populations over the past
century, but that a few lakes had exhibited changes in
sponge populations perhaps indicating a decline in water quality (Paduano and Fell, 1997). Spicule presence
and distribution has been used to infer the history of
selected Florida soils (Schwandes and Collins, 1994).
Further studies have the potential to reveal other important features of lake history.
D. Evolutionary Relationships
All freshwater sponges belong to the class Demospongiae, the most diverse and morphologically complex of the four major groups in the sponge phylum.
Within the demosponges, the evolutionary history of
the freshwater sponges is long and polyphyletic. Fossil
4. Porifera
freshwater sponges are reported at least 100 million
years before (Racek and Harrison, 1975); and, although
there is still debate in this area, there appear to have
been at least three, and possibly four, separate, successful invasions of freshwater from marine habitats. The
separate invasions are presently classified into distinct
families among the freshwater sponges (VolkmerRibeiro and De Rosa-Barbosa, 1979, Volkmer-Ribeiro,
1986).
The phylogenetic diversity of freshwater sponges
is not well represented in North America where species
are confined primarily to the most cosmopolitan
family, the Spongillidae (Penney and Racek, 1968).
Volkmer-Ribeiro (1986) proposed that species in the
genus Corvomeyenia should be grouped, along with
several South American species, in the family Metaniidae with an evolutionary history in freshwater that is
distinct from the spongillids. Only two North American species, Corvomeyenia everetti and C. carolinensis,
occur in this proposed family which has a worldwide,
primarily tropical distribution (Volkmer-Ribeiro,
1986). Not all sponge biologists have recognized the
Metaniidae as a separate family (e.g., Ricciardi and
Reiswig, 1993), however.
The two other freshwater sponge families are absent from North America. Species in the Lumbomerskiidae appear to be confined to a few large and ancient
lakes in Europe and Asia (e.g., Lakes Baikal and
Ochrid), and the family is characterized by a large degree of endemism. Species in the Potamolepidae are
widely distributed throughout Africa and South America (Volkmer-Ribeiro and De Rosa-Barbosa, 1979).
IV. COLLECTING, REARING, AND PREPARING
SPONGES FOR IDENTIFICATION
Collecting sponges is usually straightforward. In
many cases, sponges grow in shallow water and can be
obtained simply by hand or with a long-handled rake.
Where sponges are rare, it may be easiest to collect
them while snorkeling. This may be the case particularly when collecting in areas with numerous hard substrates where small stones must be overturned and the
undersides of fallen trees and large boulders must be
examined. Where sponges grow in deeper waters,
scuba techniques afford the most efficient means of collection. Dredges or nets dragged across substrates in
deeper waters are likely to miss most sponges and, at
the same time, are likely to disrupt substantial bottom
areas. In very deep waters, however, these may be the
only practical means of collecting.
When searching a lake or stream for sponges, it is
best to examine as broad a variety of substrates as
115
possible. Aquatic macrophytes, rocks, and fallen logs
are common sites for sponge growth. In bog habitats,
the roots of vegetation growing at the edges of bog
mats or on the undersides of the mats themselves are
likely sites for sponges. Where sponges are rare, it may
be necessary to examine a large number of different
substrates before finding any specimens. In other habitats, however, sponges will be conspicuous.
Growing or maintaining sponges under controlled
environmental conditions is much more difficult than
collecting them. Some investigators have grown small
sponges from gemmules in the laboratory for cytological investigations (Rasmont, 1962; Fell, 1974). Such
specimens can also be particularly useful in examining
smaller scale structural details of sponges. Poirrier et al.
(1981) have developed an effective, continuous-flow
system for sponges using bacteria as a food source.
Laboratory investigations of this sort can provide
important insights into sponge ecology (e.g., Harsha
et al., 1983), particularly when controlled conditions
are essential. Sponges appear to be highly sensitive to
environmental conditions, however, and the responses
of sponges obtained in laboratory studies must be
compared carefully with their behavior under field situations. As such, to evaluate sponge behavior under
natural conditions, it is often best to work with freshly
collected sponges and to conduct experiments in situ
whenever possible.
The identification of freshwater sponges depends
primarily on characteristics of spicules and on features
of intact gemmules. Successful species identifications
are dependent on obtaining all of the spicules (megascleres, gemmoscleres, and, if present in a species,
microscleres) that can occur in a species. Thus, it is
absolutely essential to obtain samples of sponges that
include all spicule types. Gemmoscleres are particularly
important, and this can present a problem because
gemmules (Figs. 1 and 8) may only occur during certain times of the year. Some gemmoscleres may remain
in active sponge tissue after gemmules have hatched, so
there is some chance of identifying sponges without
gemmules visible in them. Gemmules should be sought
carefully in any survey, however.
As the primary step in species identification, samples of all types of spicules that occur in a species
should be prepared for microscopic examination. To
obtain spicule preparations, organic portions of a
sponge are digested with acid and the remaining
spicules are mounted on a glass microscope slide.
Sponge tissue can be digested with nitric acid in a centrifuge tube that is immersed in boiling water for 1 h.
The spicules can then be concentrated by gentle centrifugation, after which the acid is poured off and the
spicules are then washed with ethanol or methanol.
116
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
Washing and centrifugation should be repeated at least
two times after which the spicules are mounted on a microscope slide using a cover slip and a permanent
medium (e.g., Permount). Reiswig and Browman (1987)
described a similar, but more precise technique for a
quantitative method of processing sponge spicules.
Intact gemmules can be removed from sponge tissue, dried, and mounted directly on microscope slides.
Specimens of entire sponges are necessary for museum
collections. Whole sponges can be dried or preserved in
alcohol. Simpson (1963) describes methods that can be
used for preparing freshwater sponges for cytological
observations, such as are necessary for examining reproductive cycles.
V. CLASSIFICATION
The classification scheme reported here is derived
primarily from Penney and Racek (1968), who have
completed the most recent taxonomic revision of the
family Spongillidae, including the genus Corvomeyenia.
Their monograph provides more detailed descriptions
of most of the species reported here and should be
consulted for rigorous taxonomic investigations. Information for several species in this chapter is based on
reports other than those Penney and Racek (1968),
most of which have been published since 1968. In the
cases where substantial information about a species has
been obtained from a source other than, or in addition
to, Penney and Racek, these sources have been cited
along with the species description in the list of species
below. In addition, there are a number of cases where
species reported for Canada and the United States by
Penney and Racek (1968) have been judged subsequently as invalid (e.g., Poirrier, 1974, 1977; Harrison
and Harrison, 1977; Harrison, et al., 1977; Harrison,
1979), and these species have not been included here.
Ricciardi and Reiswig (1993) provide another valuable
source of information on 15 sponge species that occur
in eastern Canada, and this report should be consulted
for more detailed information on these species.
In general, the freshwater sponge species in
Canada and the United States can be distinguished by
characteristics of their spicules. Taxa are separated
primarily by the presence, shape, and spination of microscleres and / or gemmoscleres, although the presence
or absence of spines on megascleres can be a useful
characteristic in some cases in distinguishing species
within a genus. Spicules are usually either needle or
rodlike (Fig. 6) or dumbell-shaped (Fig. 10). The latter
are termed birotulate, and their rounded ends are
called rotules. In the descriptions here, a spicule should
be assumed to be needle or rodlike, unless it is described as otherwise. Also, the descriptions reported
here are based on examinations with light microscopes.
More detailed examinations with scanning electron microscopes can reveal fine structures, primarily spines,
that are not apparent in light microscope observations.
Such finer scale observations should be interpreted with
caution when employing the descriptions and key that
are provided here.
Gemmoscleres are critical in the classification of
freshwater sponges, and it is important to include
gemmules when preparing sponge specimens for identification. Obtaining gemmoscleres may be a problem,
particularly in young sponges derived from larvae,
where gemmules and their spicules may not be present
during certain times of the year. Examine a collected
specimen closely to insure that gemmules are present
(Figs. 1 and 8). It is possible, particularly in larger specimens, that some gemmoscleres may have been left in a
sponge body from a previous year’s gemmules. However, it is also possible that a few stray gemmoscleres or
FIGURE 10 Megasclere (M, scale bar 50 m) and gemmoscleres (g, scale bar
25 m;) of Anheteromeyenia argyrosperma (from Ricciardi and Reiswig, 1993).
4. Porifera
even entire gemmules from another sponge species may
have been incorporated into a sponge specimen. The
growth of the tissue of one sponge species over the
gemmules of another species has lead to some erroneous species identifications (Ricciardi and Reiswig,
1992). Thus, identifications based on only a few gemmoscleres should be made with caution.
In some species with microscleres, it may be
difficult to distinguish between microscleres and gemmoscleres on the basis of structure alone. By separating
gemmules from intact sponges and making separate
spicule preparations of them, it is possible to obtain
samples in which gemmoscleres predominate. Alternatively, by carefully sampling tissue from the surface of a
sponge in areas of new growth it may be possible to
obtain material in which gemmoscleres are rare and in
which microscleres and megascleres predominate. Because of the important role that microscleres play in
separating species, identifications based on only a few
microscleres should also be made with caution.
It is important to consider that ecomorphic variation can be substantial for some sponge species, particularly Anheteromeyenia ryderi, Ephydatia fluviatilis, and
Spongilla lacustris. Cautions about particularly troublesome situations are raised in the species descriptions
below. Ricciardi and Reiswig (1993) provide more
detailed information on the eastern Canadian species.
In using the key presented here, there are only two
cases in which characteristics other than the structure
of spicules are required to distinguish among taxa. In
contrasting the genera Ephydatia and Radiospongilla,
it is necessary to obtain a cross section of intact gemmules in order to view the arrangement of gemmoscleres within the gemmule coat. Similarly, within the
genus Heteromeyenia, it is necessary to employ the
size, shape, and structures of the foraminal aperture of
entire gemmules to distinguish among species. Particularly important for Heteromeyenia are the forms of
structures on the end of the foraminal tubule termed
cirrous projections.
Ricciardi and Reiswig (1993) reported an extensive
survey of sponges in eastern Canada in which they
reported 15 species. The report is extremely useful and
should be consulted for detailed information on the
species that they report. It even includes a key for distinguishing, to a certain extent, among sponges for which
gemmoscleres are not available — a critical requirement
of the key here and for using Penney and Racek (1968).
A. Freshwater Sponge Species of Canada
and the United States
Summarized below are the spicular characteristics
and distribution patterns of freshwater sponges reported
117
from the United States and Canada. Dimensions presented for spicules are the ranges that have been reported for them; average values to be expected are very
close to the mean of the numbers listed below. In some
cases, mean dimension values are reported as xx(mean)-xx, where xx represents ranges. It is important
to note that these mean values are from Ricciardi and
Reiswig (1993). They may be specific to sponges from
eastern Canada and may not be representative of
sponges growing in other regions. Values for ranges are
the widest values that have been published in the papers reported here. Some sponge spicule features may
vary from habitat to habitat. For a few species information is also provided on features of gemmules, such
as the distribution of gemmoscleres within them, their
opening structures (foraminal apertures), and their
overall distribution within a sponge body. Geographic
distribution patterns emphasize the occurrence of
sponges in the United States and Canada, but also describe the overall distributions reported for each
species.
Anheteromeyenia argyrosperma (Potts)
Spicules: megascleres slender, 240-(284)-304 m in
length, and sparsely covered with small, sharply
pointed spines; microscleres absent; gemmoscleres birotulates of two distinct length groups, 65-(81)-89 or
110-(130)-160 m, with both size classes similar in
form exhibiting spines on their entire shaft and conspicuous recurved, clawlike hooks on their ends.
Distribution: reported from the eastern half of
North America from Florida to Canada but confined to
this region.
Anheteromeyenia ryderi (Potts)
Spicules: megascleres are extremely variable from
habitat to habitat with lengths ranging from 141-(220)279 m; variable shape and coverage with broadly
conical spines; microscleres absent; gemmoscleres birotulates of two distinct length groups with distinct differences in their shapes, 28-(34)-41 or 45-(64)-75 m
in length; shorter forms have shafts with only one or a
few spines and flattened rotules with a large number of
small teeth; larger forms are robust with numerous recurved spines on their shaft and with strongly recurved
hooks on their ends.
Distribution: reported from the eastern half of
North American, from Louisiana to Canada, but confined to this region.
Corvomeyenia carolinensis (Harrison)
Spicules: megascleres slender, straight to slightly
curved and entirely smooth, ranging 194 – 280 m in
length; microscleres birotulate with straight to strongly
118
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
FIGURE 11 Megasclere (M, scale bar 50 m) and gemmoscleres (g, scale bar
25 m;) including a rotule (r) of Anheteromeyenia ryderi (from Ricciardi and Reiswig,
1993).
curved ( 80%), smooth shafts, 15 – 25 m in length
terminating in rotules 4 – 7 m in diameter with 4 – 6
recurved hooks; gemmoscleres birotulates with straight
to slightly curved, smooth shafts 60 to 158 m in
length and terminating in rotules of 13 – 22 m diameter with 5 – 8 recurved hooks (Harrison, 1971).
Distribution: reported from one pond in South
Carolina and one lake in Connecticut.
gemmoscleres birotulate with straight to slightly
curved, smooth shafts, 33-(59)-78 m in length and
terminating in rotules 10-(20)-26 m in diameter
bearing 5 – 7 recurved hooks. See information on
Corvospongilla novaeterrae below and in Ricciardi and
Reiswig (1993).
Distribution: reported only from the eastern half of
the northern United States and Canada.
Corvomeyenia everetti (Mills)
Spicules: megascleres slender, slightly curved and
entirely smooth, 143-(218)-285 m in length (in rare
cases, a variable number of megascleres may be
sparsely spined); microscleres birotulates, 14-(18)26 m in length, terminating in rotules 3-(5)-7 m in
diameter with 3 – 6 small, distinctly recurved spines;
Corvospongilla becki (Poirrier)
Spicules: megascleres stout, 130 – 218 m in length,
usually curved, covered with spines, the spines being
larger near the ends of a spicule; microscleres birotulate,
25 – 44 m in length, rotules 9 – 17 m in diameter, usually with four recurved hooks; gemmoscleres of two distinct length classes, the smaller class is 28 – 56 m in
length, slightly to strongly curved and spined, except in
the inner curved region, the larger class 71 – 139 m in
length, straight to slightly curved and completely spined
(somewhat similar in form to the gemmoscleres of S. lacustris — Fig. 29) (Poirrier, 1978). See information on
Corvospongilla novaeterrae below.
Distribution: reported from one lake in Louisiana.
FIGURE 12
Microscleres (m) and gemmoscleres (G) of Corvomeyenia carolinensis (after Harrison, 1971).
Corvospongilla novaeterrae (Potts)
Spicules: megascleres stout, smooth, 112-(154)-170
m in length and relatively scarce; microscleres are
abundant birotulates, 13-(21)-32 m in length with
smooth shafts, rotules dome shaped with 3 – 6 spines;
gemmoscleres highly variable, bearing numerous large,
recurved spines which tend to be concentrated near the
ends of the shaft, sometimes making spicules appear
birotulate with lengths 21-(39)-63 m and widths, excluding spines, 3-(6)-9 m. This species is remarkably
similar to Corvomeyenia everetti in its basic growth
form, except that C. novaeterrae gemmoscleres are not
thin, elongate birotulates and C. novaeterrae gemmules
4. Porifera
119
FIGURE 13 Megasclere (M, scale bar 50 m), microscleres (m, scale bar 25 m)
including a rotule (r) and gemmosclere (g, scale bar 25 m) of Corvomeyenia everetti
(from Ricciardi and Reiswig, 1993).
have a very poorly developed outer layer. See detailed
information on C. novaeterrae in Ricciardi and Reiswig
(1993).
Distribution: reported from a few lakes in the maritime provinces of Canada and one lake in Connecticut
(De Santo and Fell, 1996).
Dosilia palmeri (Potts)
Spicules: megascleres slender, 370 – 450 m in
length, slightly curved to nearly straight, covered with
sparse spines in their central portion; microscleres stellate with 8 – 12 usually smooth rays arising from a
central nodule, length is extremely variable; gemmoscleres birotulates, 55 – 85 m in length, occurring
in two subtly distinct size classes, with strong spines on
their central shaft and with equally sized rotules,
23 – 25 m in diameter, bearing numerous blunt recurved teeth (Penney, 1960).
Distribution: reported from Florida in North
America north of Mexico and from other locations in
Central America.
Dosilia radiospiculata (Mills)
Spicules: megascleres slender, 290 – 400 m in
length, entirely smooth or covered with minute spines;
microscleres stellate with 6 – 8 microspined rays projecting from their center, length is extremely variable;
gemmoscleres birotulates of two distinctly different size
classes with longer forms exhibiting nonspined shafts
120 – 230 m in length and shorter forms exhibiting
strongly spined shafts 45 – 82 m in length. [There is
some question as to whether the two species of Dosilia
are distinct or simply ecomorphic variants of a single
species.]
Distribution: reported from the Canadian border
south to Mexico, but only from this region.
Megasclere (M, scale bar 50 m), microscleres (m, scale bar
10 m) and gemmoscleres (g, scale bar 10 m) of Corvospongilla novaeterrae (from
Ricciardi and Reiswig, 1993).
FIGURE 14
120
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
FIGURE 15
Microscleres of Dosilia radiospiculata (after Penney
and Racek, 1968).
Duosclera mackayi (Carter)
Spicules: two distinct classes of megascleres, the
first is relatively scarce, straight or slightly curved, 177(200)-302 m in length, 7-(12)-18 m in width
(excluding spines), covered with coarse spines and the
second is somewhat shorter, 79-(156)-267 m in length
and generally broader, 2-(8)-20 m in width (excluding
spines) and densely covered with spines that are
long, pointed, strongly recurved near the tips of the
spicule and perpendicular near its center; microscleres
absent; gemmoscleres are the same as the second class
of megascleres, which are always present in a specimen,
even in the absence of gemmules. When intact
gemmules are present, gemmoscleres are arrayed
tangentially to the surface of the gemmule except near
the foraminal aperture. The overall orientation of
gemmules can help distinguish Duosclera mackayi
from a similar species, E. fragilis. In D. Mackayi
the foraminal apertures are always oriented inward
or towards a substrate while those in E. fragilis are
always directed outward or upward from a pavement
layers. This species has recently been classified as a new
genus (Reiswig and Ricciardi, 1993). Previously reported as Eunapius igloviformis in Penney and Racek
(1968) and as Eunapius mackayi in Ricciardi and
Reiswig (1993).
Distribution: throughout the United States and
Canada, but confined to these regions.
Ephydatia fluviatilis (Linneaus)
Spicules: megascleres usually slightly curved, 210(343)-439 m in length, and usually entirely smooth,
although in some cases, some sparsely spined megascleres co-occur with smooth forms; microscleres absent;
gemmoscleres birotulates of one class, 20-(23)-30 m
in length with a slender, smooth shaft, which sometimes has 1 – 4 large spines, with flat irregularly shaped
rotules of equal, 13-(18)-24 m diameters and usually
more than 20 teeth that are not deeply incised. E.
fluviatilis can sometimes be confused with E. muelleri.
These species can be most surely distinguished by
comparing gemmosclere length to rotule diameter.
Megascleres (M, scale bars 50 and 25 m for the magnified section),
gemmoscleres, which are actually a second size category of megasclere (g, scale bars
25 m) and gemmules of Duosclera mackayi, the arrowhead indicates the foraminal
aperture on the underside of a gemmule cluster (from Ricciardi and Reiswig, 1993).
FIGURE 16
4. Porifera
121
Megasclere (M, scale bar 50 m) and gemmoscleres (g, scale bar 25
m) and a rotule (r, scale bar 10 m) of Ephydatia fluviatilis (from Ricciardi and
Reiswig, 1993).
FIGURE 17
Gemmosclere length is always greater than rotule diameter in E. fluviatilis while gemmosclere length is less
than or equal to rotule diameter in E. muelleri (Ricciardi and Reiswig, 1993).
Distribution: appears to be truly cosmopolitan
with more frequent occurrence in temperate than in
tropical zones.
Ephydatia millsii (Potts)
Spicules: megascleres slightly curved to nearly
straight, 180 – 270 m in length, with numerous small
spines except at the tips; microscleres absent; gemmoscleres birotulates of one size class, 36 – 48 m in
length, with smooth shafts that are clearly broader near
the rotules and with distinctly flat, disk – shaped rotules
of equal, 22 – 28 m diameters and only very small incisions at their margins.
Distribution: reported only from Florida.
Ephydatia muelleri (Lieberkühn)
Spicules: megascleres straight to slightly curved,
171-(245)-350 m in length, usually with small spines,
typically lacking at the tips, but smooth in rare cases;
FIGURE 18
Racek, 1968).
Gemmoscleres of Ephydatia millsii (after Penney and
microscleres absent; gemmoscleres birotulate of one
class, 8-(17)-28 m in length, with thick smooth shafts
and with flat irregularly shaped rotules of equal,
8-(15)-27 m diameters and usually have fewer
than 12 teeth that are deeply incised into long rays. E.
muelleri can sometimes be confused with E. fluviatilis.
These species can be most surely distinguished by
comparing gemmosclere length to rotule diameter.
Gemmosclere length is always greater than rotule
diameter in E. fluviatilis while gemmosclere length is
less than or equal to rotule diameter in E. muelleri
(Ricciardi and Reiswig, 1993).
Distribution: widely distributed throughout the
Northern Hemisphere with a preference for temperate
regions.
Eunapius fragilis (Leidy)
Spicules: megascleres entirely smooth, 165-(189)271 m in length; microscleres absent; gemmoscleres
straight to slightly curved, covered with conspicuous
spines often more dense near the tips, 32-(57)-140 m.
Mature gemmules enclosed in a common brown coat
forming a pavement layer cemented to the substrate
(Gp in Fig. 20) or in individual clusters of 2 – 4 gemmules. The overall orientation of gemmules can help
distinguish E. fragilis from a similar species, Duosclera
mackayi. In E. fragilis, the foraminal apertures are
always directed outward from a cluster or upward
from a pavement layer while those in D. mackayi
are always oriented inward or toward a substrate
(Ricciardi and Reiswig, 1993).
Distribution: truly cosmopolitan.
Heteromeyenia baileyi (Bowerbank)
Spicules: megascleres slender, 216-(247)-320 m in
length, smooth or with sparse microspines except near
the tips; microscleres 53-(67)-85 m in length, delicate,
122
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
Spined and smooth megascleres (M, scale bar 50 m) and gemmoscleres (g, scale bar 10 m), and a rotule (r, scale bar 10 m) of Ephydatia
mülleri (from Ricciardi and Reiswig, 1993).
FIGURE 19
slightly curved to almost straight with spines that occur
throughout their length, but which increase in length
towards the central region where they are often
knobbed and distinctly perpendicular to the main axis;
gemmoscleres birotulate of two intergrading length
classes: (1) short birotules with flat, serrated rotules
(13-(18)-22 m in diameter), 38-(51)-60 m in length,
and (2) long birotules 49-(70)-86 m in length with
rotules (18-(22)-28 m in diameter) composed of long
recurved hooks giving an umbrellalike appearance
(hooks often have knobbed tips). Foraminal apertures
of gemmules do not have terminal cirrous projections
like those for H. tubisperma (Fig. 24), H. latitenta
(Fig. 22), and H. tentasperma (Fig. 23). The long
Megasclere (M, scale bar 50 m), gemmoscleres (g, scale bars
25 m), and a gemmule pavement of Eunapius fragilis, the arrowhead indicates a
foraminal aperture on the dorsal surface of the gemmule pavement (from Ricciardi and
Reiswig, 1993).
FIGURE 20
4. Porifera
123
FIGURE 21 Megasclere (M, scale bar 50 m), microsclere (m, scale bar 25 m)
and gemmoscleres (g, scale bar 25 m) of Heteromeyenia baileyi (from Ricciardi and
Reiswig, 1993).
middle spines on the microscleres of H. baileyi are also
useful in distinguishing it from other species in the
genus. Their maximum length is on average greater
than the width of the microsclere.
Distribution: widely distributed throughout the
eastern United States and Canada with a few reports
from Europe.
Heteromeyenia latitenta (Potts)
Spicules: megascleres straight, 265 – 285 m in
length, smooth to sparsely microspined; microscleres
slender and entirely spined with those in the central region only slightly larger than those on the ends,
85 – 100 m; gemmoscleres birotulates of two length
groups, 50 – 55 or 60 – 78 m with shafts bearing numerous stout and pointed spines, both rotules of equal,
16 – 18 m diameters, margins forming numerous conspicuous, recurved teeth. Foraminal apertures of gemmules with one or two, very long, cirrous projections
starting from a disk that is initially flat but rounded in
its terminal regions.
FIGURE 22
Foraminal aperture of Heteromeyenia latitenta (after
Neidhoefer, 1940).
Distribution: reported only from northeastern
United States.
Heteromeyenia tentasperma (Potts)
Spicules: megascleres very slender, 260 – 280 m in
length, with sparse microspines; microscleres slender
with sparse microspines, 75 – 80 m; gemmoscleres birotulates of two length groups, 50 – 55 or 65 – 72 m with
stout shafts bearing a small number of acute spines and
with both rotules of equal, 15-18 m diameters and
consisting of an arrangement of lateral spines. Foraminal
apertures of gemmules distinctly tubular and relatively
short with 3 – 6 long and irregular cirrous projections.
Distribution: reported only from northeastern to
midwestern United States.
Heteromeyenia tubisperma (Potts)
Spicules: megascleres slender, 190-(290)-337 m in
length, smooth to sparsely microspined; microscleres,
73-(100)-118 m in length, slender and entirely spined
with spines near the tips small and recurved and those
near the central portion distinctly larger, straight, and
with knobs; gemmoscleres birotulates of two roughly
defined size classes, with both classes 33-(44)-70 m in
length with stout shafts bearing a small number of acute
spines and with both rotules of equal, 12-(19)-25 m
FIGURE 23
Foraminal aperture of Heteromeyenia tentasperma
(after Neidhoefer, 1940).
124
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
FIGURE 24 Megasclere (M, scale bar 50 m), microsclere (m, scale bar 25 m),
and gemmoscleres (g, scale bar 25 m) and a foraminal aperture (GF) of
Heteromeyenia tubisperma (from Ricciardi and Reiswig, 1993).
diameters, consisting of an arrangement of lateral
spines. Foraminal apertures of gemmules distinctly
tubular, slender and very long (0.5 – 0.9 times the
diameter of the gemmule) with 5 or 6 cylindrical cirrous
projections. It is important to note that developing gemmules may have stunted foraminal development such
that they resemble H. latitenta and H. tentasperma.
Specimens should be thoroughly inspected for fully developed gemmules. H. tentasperma can be distinguished
from H. baileyi which has shorter microscleres with
longer spines.
Distribution: reported only from the eastern half of
North America.
Radiospongilla cerebellata (Bowerbank)
Spicules: megascleres straight to slightly curved,
240 – 330 m in length, entirely without spines; microscleres absent although immature gemmoscleres
may be abundant in some portions of the dermal
membrane; gemmoscleres usually distinctly curved and
rarely straight, 72 – 110 m in length, covered with
abundant spines that are pronouncedly recurved
toward the terminal ends (in intact gemmules, gemmoscleres are arrayed in two distinct layers with those
in the inner layer arranged in a radial fashion and
those in the outer layer arranged tangentially to the
inner layer) (Poirrier, 1972).
FIGURE 25 Megasclere (M, scale bar 50 m), gemmoscleres (g, scale bar 25 m),
and a gemmule (G) showing a crater like depression around the foraminal aperture of
Radiospongilla crateriformis (from Ricciardi and Reiswig, 1993).
4. Porifera
125
Distribution: reported only from Texas in the
United States, but widely distributed in tropical and
subtropical Asia and Africa.
Radiospongilla crateriformis (Potts)
Spicules: megascleres slender and slightly curved,
240-(278)-300 m in length, sparsely covered by very
small spines, except at their tips; microscleres absent;
gemmoscleres slender and covered with small conical
spines, except at the tips where slightly recurved spines
are sufficiently dense to form pseudorotules, 60-(71)80 m in length. Gemmoscleres in intact gemmules are
arranged radially around the gemmule coat except in
the immediate vicinity of the foraminal aperture where
they form a craterlike depression leaning away from
the aperture.
Distribution: reported primarily from the eastern
half of the United States but as far west as Wisconsin
and Texas; eastern Canada (Ricciardi and Reiswig,
1993), also reported in China, Japan, Southeast Asia,
and Australia.
Spongilla alba (Carter)
Spicules: megascleres entirely smooth, 144 – 420 m
in length; microscleres slender and slightly curved
with erect spines that are longer in the central region, 49 – 124 m; gemmoscleres slightly to moderately
curved with stout, sharp, recurved spines that are more
dense at the ends, forming distinct heads, 48 – 130 m
(Poirrier, 1976).
Distribution: occurs in warmer regions worldwide with a strong preference for brackish water, re-
FIGURE 26 Microsclere (m) and gemmosclere (G) of Spongilla
alba (after Penney and Racek, 1968).
ported from the southeast coastal regions of the United
States.
Spongilla aspinosa (Potts)
Spicules: megascleres slender and entirely smooth,
155-(274)-338 m in length; microscleres range from
rare to abundant in number, smooth or very sparsely
microspined, 21-(50)-78 m in length; gemmoscleres
smooth, resembling small megascleres, 129-(274)306 m in length.
Distribution: reported from the United States from
Florida to Michigan, and from eastern Canada.
Spongilla cenota (Penney and Racek)
Spicules: megascleres stout and completely smooth,
310 – 410 m in length; microscleres numerous and
FIGURE 27 Megasclere (M, scale bar 50 m), microscleres showing a range of
forms (m, scale bar 25 m) and gemmosclere (g, scale bar 25 m) of Spongilla
aspinosa (from Ricciardi and Reiswig, 1993).
126
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
Spongilla lacustris (Linneaus)
Spicules: megascleres entirely smooth, 158-(254)362 m in length; microscleres densely covered with
small spines, 30-(61)-130 m in length; gemmoscleres
absent from winter gemmules and slightly curved, usually covered with strong, curved spines that tend to be
concentrated near the tips, 18-(32)-130 m long in
thick-coated summer gemmules (Poirrier et al., 1987).
Distribution: throughout the Northern Hemisphere
including frequent reports from throughout the United
States and Canada.
FIGURE 28 Microsclere (m) and gemmosclere (G) of Spongilla
cenota (after Penney and Racek, 1968).
slender, covered with small spines at their tip and
with a group of clearly larger spines in their center,
68 – 123 m; gemmoscleres extremely stout, entirely
covered with stout, sharp, recurved spines 65 – 86 m
(Poirrier, 1976).
Distribution: reported only from Florida, and the
Yucatan in Mexico.
Stratospongilla penneyi (Harrison)
Spicules: megascleres slightly curved, 215 – 296 m
in length, smooth to very delicately microspined, microscleres slender, 38 – 75 m in length, covered with
very small spines, gemmoscleres curved to distinctly
bent, 48 – 123 m in length, smooth to delicately microspined with sharply pointed apices.
Distribution: reported from only one location, a
canal in southern Florida (Harrison, 1979).
Trochospongilla horrida (Weltner)
Spicules: megascleres straight to slightly curved,
155-(187)-250 m in length, covered with stout,
FIGURE 29 Megasclere (M, scale bar 50 m), microscleres showing a range of aberrant forms in the right plate (m, scale bar 25 m) and gemmosclere (g, scale bar 25
m) of Spongilla lacustris (from Ricciardi and Reiswig, 1993).
4. Porifera
127
Megascleres (M, scale bars 50 and 25 m for magnified portion which
shows the truncated spines typical of the genus) and gemmoscleres (g, scale bar 10
m) with rotule (r) of Trochospongilla horrida (from Ricciardi and Reiswig, 1993).
FIGURE 30
blunt, truncated spines; microscleres absent; gemmoscleres small birotulates, 8 – 10 m in length, with
stout smooth shafts and with rotules of nearly equal
size, with the smaller between 9 and 13 m in diameter and the larger between 13 and 16 m with circular margins (overall length of gemmoscleres is never
greater than the diameter of the smaller rotule). Compare with information for T. pennsylvanica to confirm
an identification.
Distribution: widely dispersed throughout the
Northern Hemisphere although with few reports in
eastern Canada.
to this chapter. In such cases, the two species can be
distinguished by the length of the gemmosclere shaft
relative to the rotule diameter. The shaft length is
longer than, or in rare cases, equal to, the diameter for
the smaller rotule’s diameter in T. pennsylvanica and
shorter than the rotule’s diameter for T. horrida
(Ricciardi and Reiswig, 1993).
Distribution: reported throughout, but apparently
restricted to, the North American continent.
Trochospongilla leidii (Bowerbank)
Spicules: megascleres straight to slightly curved
and entirely smooth, 150 – 170 m in length; microscleres absent; gemmoscleres small birotulates, 11
m in length, terminating in rotules with circular margins and equal, 12 – 14 m diameters.
Distribution: reported from limited regions of the
eastern United States, with one report of a population
from the Panama Canal (Jones and Rützler, 1975).
Please note that the successful use of this key depends on obtaining a fully representative sample of all
Trochospongilla pennsylvanica (Potts)
Spicules: megascleres slender and slightly curved,
100-(253)-432 m in length, entirely covered with
blunt, truncated spines; microscleres absent; gemmoscleres small birotulates with slender shafts, 11(17)-41 m in length, terminating in rotules with circular margins and usually with two distinctly different
diameters, 3-(9)-23 m and 13-(24)-41 m. In some
cases both rotules of the gemmoscleres of T. pennsylvanica appear to have the same diameters leading them
to be identified incorrectly as T. horrida using the key
B. Key to the Freshwater Sponges in Canada and the
United States
FIGURE 31 Megasclere (M) and gemmosclere (G) of Trochospongilla
leidii (after Penney and Racek, 1968).
128
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
Megasclere (M, scale bars 50 m) and gemmoscleres (g, scale bar
10 m) with rotules (r) or aberrant forms (ga) of Trochospongilla pennsylvanica (from
Ricciardi and Reiswig, 1993).
FIGURE 32
of the types of spicules that can occur within a
sponge species (megascleres, gemmoscleres, and, if
they occur in a species, microscleres). If a species ordinarily exhibits gemmoscleres or microscleres and
they are not contained within a specimen, it will not
be possible to identify even its genus. The one exception to this rule occurs for Spongilla lacustris which
is unique among the species in this key in its lack of
gemmoscleres in its winter gemmules. If gemmules
are clearly present in a specimen but gemmoscleres
are absent, it is most likely S. lacustris. [Caution is
necessary here in that some foreign bodies in sponges
may resemble gemmules superficially, for example,
the eggs of water mites. Gemmules can be distinguished by their highly resistant coats.] It is important to note that S. lacustris. may also have gem-
moscleres; they occur when summer gemmules are or
have been present.
Also note that this key is intended only for use with
the species listed above from Canada and the United
States. The couplets in this key have been designed only
for this particular subset of the world’s freshwater
sponges. Those examining specimens from other regions
will need to consult Penney and Racek (1968) and more
recent taxonomic work (e.g., Volkmer-Ribiero, 1986, and
the references cited in the species list). The key is arranged
systematically so that genera are separated prior to species
within a genus. In most cases, therefore, a valid identification to genus may be possible even if a species has not
previously been reported in Canada or the United States.
Even to separate genera, however, it is necessary to have
included all the possible spicule types from a specimen.
1a.
Microscleres present. ..........................................................................................................................................................................2
1b.
Microscleres absent. .........................................................................................................................................................................16
2a(1a).
Microscleres star-shaped (Fig. 15) or
birotulate (Figs. 10 and 11). ...............................................................................................................................................................3
2b.
Microscleres rod-shaped to needlelike in structure.
(Fig. 21 and Fig. 24)...........................................................................................................................................................................8
3a(2a).
Microscleres star-shaped with several rays extending from central region of the spicules
(Fig. 15)......................................................................................................................................................................Genus Dosilia 4
3b.
Microscleres birotulate (Figs. 12 and 13) ...........................................................................................................................................5
4a(3a).
Gemmoscleres birotulate in two distinctly different size categories (45 – 82 m in length and 120 – 230 m in length).
..........................................................................................................................................................................Dosilia radiospiculata
4b.
Gemmoscleres birotulate in two size categories that are nearly equal in length (55 – 85 mm).
.....................................................................................................................................................................................Dosilia palmeri
5a(3b).
Gemmoscleres birotulate (Figs. 12 and 13).
.......................................................................................................................................................................Genus Corvomeyenia 6
4. Porifera
5b
129
Gemmoscleres rod- or needle-shaped (Fig. 14).
.......................................................................................................................................................................Genus Corvospongilla 7
6a(5a).
Microscleres straight to slightly curved birotules with a predominance of straight forms (Fig. 12).
..........................................................................................................................................................................Corvomeyenia everetti
6b.
Microscleres a mixture of straight to strongly curved birotules with a predominance of curved forms (Fig. 11).
..................................................................................................................................................................Corvomeyenia carolinensis.
7a(5b).
Megascleres covered with spines
............................................................................................................................................................................Corvospongilla becki
7b.
Megascleres smooth
..................................................................................................................................................................Corvospongilla novaeterrae
8a(2b).
Gemmoscleres birotulate (as in Fig. 21).
.......................................................................................................................................................................Genus Heteromeyenia 9
8b.
Gemmoscleres needlelike (as in Fig. 17) or absent. ...........................................................................................................................12
9a(8a).
Foraminal aperture of gemmules without terminal cirrous projections (contrast with Figs. 22, 23, and 24).
..........................................................................................................................................................................Heteromeyenia baileyi
9b.
Foraminal aperture of gemmules with distinct, terminal cirrous projections (Figs. 22, 23, and 24). .................................................10
10a(9b).
Foraminal aperture of gemmules with one to two very long cirrous projections starting from a flat disk, flat and ribbonlike at the
base and cylindrical at the end. (Fig. 22) .........................................................................................................Heteromeyenia latitenta
10b.
Foraminal aperture of gemmules with 3 – 6 cirrous projections that are short when compared with those of H. latitenta. ...............11
11a(10b).
Foraminal aperture of gemmules distinctly tubular and very long ranging from 0.5 to 0.9 times the diameter of the gemmule
(Fig. 23) .....................................................................................................................................................Heteromeyenia tubisperma
11b.
Foraminal aperture of gemmules distinctly tubular, but short, less than 0.4 times the diameter of the gemmule (Fig. 24).
.................................................................................................................................................................Heteromeyenia tentasperma
12a(8b).
Gemmoscleres smooth or covered with very fine spines....................................................................................................................13
12b.
Gemmoscleres absent or covered with robust, conspicuous spines (Figs. 28 and 29).
.............................................................................................................................Genus Spongilla, except for S. aspinosa ..............14
13a(12a).
Microscleres smooth or very sparsely microspined (Fig. 27). ...................................................................................Spongilla aspinosa
13b.
Microscleres covered with very small spines. ...................................................................................................Stratospongilla penneyi
14a(12a).
Gemmoscleres clublike with spines conspicuously more dense on the ends than in the center (Fig. 26). .........................Spongilla alba
14b.
Gemmoscleres absent or with spines distributed evenly across the length of the spicule (Figs. 28 and 29). .......................................15
15a(14b).
Microscleres with conspicuously denser and longer spines in the central region (Fig. 28). ..........................................Spongilla cenota
15b.
Microscleres with spines distributed evenly across the length of the spicule (Fig. 29). ..............................................Spongilla lacustris
16a(1b).
Megascleres of a single size category with distinctly different forms of gemmoscleres ......................................................................17
16b.
Megascleres in two distinctly different size categories (Fig. 16) with the smaller category serving also as gemmoscleres although
both sizes of megascleres are always present even when gemmules are absent. .......................................................Duosclera mackayi
17a(16a).
Gemmoscleres birotulate (Fig. 30) ....................................................................................................................................................18
17b.
Gemmoscleres rod shaped or needlelike (Fig. 20) .............................................................................................................................25
18a(17a).
Margins of rotules completely smooth (Figs. 30 and 31).
....................................................................................................................................................................Genus Trochospongilla 19
18b.
Margins of rotules distinctly spined or serrated (Figs. 17 and 19) ....................................................................................................21
19a(18a).
Megascleres conspicuously spined (Fig. 30) ......................................................................................................................................20
19b.
Megascleres smooth or with very fine spines (Fig. 31) ........................................................................................Trochospongilla leidii
130
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
20a(19a).
Gemmoscleres with rotules of two distinctly different diameters (Fig. 32) ............................................Trochospongilla pennsylvanica
20b.
Gemmoscleres with rotules of nearly equivalent diameters. (Fig. 30) ..............................................................Trochospongilla horrida
21a(18b).
Gemmoscleres occurring in two distinct size classes (Fig. 10)
..................................................................................................................................................................Genus Anheteromeyenia 22
21b.
Gemmoscleres occurring in only one size class (Fig. 17)
.............................................................................................................................................................................Genus Ephydatia 23
22a(21a).
Gemmoscleres of two classes that are similar in shape, but clearly different in length (Fig. 10)
...........................................................................................................................................................Anheteromeyenia argyrosperma
22b.
Gemmoscleres of two classes that are distinct in both shape and size (Fig. 11).
.......................................................................................................................................................................Anheteromeyenia ryderi
23a(21b).
Rotules of gemmoscleres with margins that are nearly smooth bearing numerous, very small incisions (Fig. 18).
..................................................................................................................................................................................Ephydatia millsii
23b.
Rotules of gemmoscleres with clearly indented margins (Fig. 17). ....................................................................................................24
24a(23b).
Gemmoscleres less than 20 m in length with rotules bearing fewer that 12 teeth deeply incised into long rays, gemmosclere length
less than or equal to rotule diameter (Fig. 19).
...............................................................................................................................................................................Ephydatia muelleri
24b.
Gemmoscleres greater than 25 m in length with rotules bearing greater than 20 teeth that are not deeply incised (Fig. 17).
.............................................................................................................................................................................Ephydatia fluviatilis
25a(17b).
Most gemmoscleres in intact gemmules arrayed in a distinctly radial fashion within the gemmule coat (Fig. 25).
.....................................................................................................................................................................Genus Radiospongilla 26
25b.
Gemmoscleres in intact gemmules arrayed only tangentially to the surface of the gemmule.
..................................................................................................................................................................................Eunapius fragilis
26a(25a).
Megascleres sparsely covered with small spines (Fig. 25).
................................................................................................................................................................Radiospongilla crateriformis
26b.
Megascleres entirely smooth.
....................................................................................................................................................................Radiospongilla cerebellata
LITERATURE CITED
Bader, R. B. 1984. Factors affecting the distribution of a freshwater
sponge. Freshwater Invertebrate Biology 3:86 – 95.
Barbeau, M. A., Reiswig, H. M., Rath, L. C. 1989. Hatching of
freshwater sponge gemmules after low temperature exposure:
Ephydatia mülleri (Porifera: Spongillidae). Journal of Thermal
Biology 14:225 – 231.
Barton, S. H., Addis, J. S. 1997. Freshwater sponges (Porifera:
Spongillidae) of western Montana. Great Basin Naturalist
57:93 – 103.
Bergquist, P. R. 1978. Sponges, University of California Press, Berkeley, CA.
Berzins, B. 1950. Observations on rotifers on sponges. Transactions
of the American Microscopical Society 69:189 – 193.
Bisbee, J. W. 1992. Life cycle, reproduction and ecology of freshwater sponges in a South Carolina pond. I. Life cycle and
reproduction of Spongilla lacustris. Transactions of the
American Microscopical Society 111:77 – 88.
Boury-Esnault, N., Rützler, K. 1997. Thesaurus of sponge morphology, Smithsonian Contributions to Zoology Number 596,
Smithsonian Institution Press, Washington, DC.
Cecil, J. T., Stempien, M. F. Jr., Ruggieri, G. D. Nigrelli, R. F. 1976.
Cytological abnormalities in a variety of normal and transformed cell lines treated with extracts from sponges, in: Harrison, F. W., Cowden, R. R. Eds., Aspects of sponge biology, Academic Press, New York, NY, pp. 171 – 182.
Clifford, H. F. 1991. Aquatic invertebrates of Alberta, The University
of Alberta Press, Edmonton, Alberta, Canada.
Colby, A. C. C., Frost, T. M., Fischer, J. M. 1999. Sponge distribution and lake chemistry in northern Wisconsin lakes: Minna
Jewell’s survey revisited. Memoirs of the Queensland Museum
44:93–99.
De Santo, E. M., Fell, P. E. 1996. Distribution and ecology of freshwater sponges in Connecticut. Hydrobiologia 341:81 – 89.
De Vos, L., Rützler, K., Boury-Esnault, N., Donadey C., Vacelet. J.
1991. Atlas of sponge morphlogy, Smithsonian Institution Press,
Washington, DC.
Dominey, W. 1987. Sponge-eating by Pungu maclareni, an endemic
Cichlid fish from Lake Barombi Mbo, Cameroon. National Geographic Research 3:389 – 393.
Fell, P. E. 1974. Porifera, in: Giese, A. C., Pearse, J. S. Eds., Reproduction of marine invertebrates, Vol. I, Academic Press, New
York, NY, pp. 51 – 132.
4. Porifera
Frey, D. G. 1963. Limnology in North America. University of Wisconsin Press, Madison, WI.
Frost, T. M. 1976. Investigations of the aufwuchs of freshwater
sponges. I. A quantitative comparison between the surfaces of
Spongilla lacustris and three aquatic macrophytes. Hydrobiologia 50:145 – 149.
Frost, T. M. 1976. Sponge feeding: A review with a discussion of
some continuing research, in: Harrison, F. W., Cowden, R. R.
Eds., Aspects of sponge biology, Academic Press, New York,
NY, pp. 283 – 298.
Frost, T. M. 1978. The impact of the freshwater sponge Spongilla lacustris on a sphagnum bog pond. Verhandlungen Internationale
Vereinigung für Theoretische und Angewandte Limnologie
20:2368 – 2371.
Frost, T. M. 1980a. Clearance rate determinations for the freshwater
sponge Spongila lacustris: effect of temperature, particle type
and concentration, and sponge size. Archiv für Hydrobiologie
90:330 – 356.
Frost, T. M. 1980b. Selection in sponge feeding processes, in: Smith,
D. C., Tiffon, Y. Eds., Nutrition in lower Metazoa, Pergammon
Press, Oxford, England, pp. 33 – 44.
Frost, T. M. 1981. Analysis of ingested materials within a freshwater
sponge. Transaction of the American Microscopical Society
100:271 – 277.
Frost, T. M. 1987. Porifera in: Pandian, T. J., Vernberg, J. F. Eds., Animal energetics, Vol. 1, Academic Press, New York, NY, pp. 27–53.
Frost, T. M., de Nagy, G. S., Gilbert, J. J. 1982. Population dynamics
and standing biomass of the freshwater sponge, Spongilla lacustris. Ecology 63:1203 – 1210.
Frost, T. M., Elias, J. E. 1990. The balance of autotrophy and heterotrophy in three freshwater sponges with algal symbionts, in:
Rützler, K. Ed., Proceedings of the Third International Conference on Sponge Biology, Smithsonian Press, Washington, DC,
pp. 478 – 484.
Frost, T. M., Graham, L. E., Elias, J. E., Haase, M. J., Kretchmer,
D. W., Kranzfelder, J. A. 1997. A yellow-green algal symbiont
in the freshwater sponge Corvomeyenia everetti: convergent evolution of symbiotic associations. Freshwater Biology 38:395 – 399.
Frost, T. M., Williamson, C. E. 1980. In situ determination of the effect of symbiotic algae on the growth of the freshwater sponge
Spongilla lacustris. Ecology 61:1361 – 1370.
Gilbert, J. J. 1974. Field experiments on sexuality in the freshwater
sponge Spongilla lacustris. Control of oocyte production and
the fate of unfertilized oocytes. The Journal of Experimental
Zoology 188:165 – 178.
Gilbert, J. J. 1975. Field experiments on gemmulation in the freshwater sponge Spongilla lacustris. Transactions of the American Microscopical Society 94:347 – 356.
Gilbert, J. J., Allen, H. L. 1973a. Studies on the physiology of the
green freshwater sponge Spongill lacustris: primary productivity, organic matter, and chlorophyll content. Verhandlungen Internationale Vereinigung für Theoretische und Angewandte
Limnlogie 20:2368 – 2371.
Gilbert, J. J., Allen, H. L. 1973b. Chlorophyll and primary productivity of some green, freshwater sponges. Internationale Revue
der Gesamten Hydrobiologie 58:633 – 658.
Gilbert, J. J., Simpson, T. L. 1976a. Sex reversal in a freshwater
sponge. The Journal of Experimental Zoology 195:145 – 151.
Gilbert, J. J., Simpson, T. L. 1976b. Gemmule polymorphism in the
freshwater sponge Spongilla lacustris. Archiv für Hydrobiologie
78:268 – 277.
Gregson, R. P., Baldo, B. A., Thomas, P. G., Quinn, R. J., Berquist, P.
R., Stephens, J. F., Horne, A. R., 1979. Flourine is a major constituent of the marine sponge Halichondria moorei. Science
206:1108 – 1109.
131
Harrison, F. W. 197l. A taxonomical investigation of the genus Corvomeyenia Weltner (Spongillidae) with an introduction of
Corvomeyenia carolinensis sp. nov. Hydrobiologia 38:123 – 140.
Harrison, F. W. 1974. Sponges (Porifera: Spongillidae). in: Hart, C.
W. Jr., Fuller, S. L. H. Eds., Pollution ecology of freshwater invertebrates, Academic Press, New York, NY, pp. 29 – 66.
Harrison, F. W. 1979. The taxonomic and ecological status of the environmentally restricted spongillid species of North America. V.
Ephydatia subtilis (Weltner) and Stratospongilla penneyi sp.
nov. Hydrobiologia 62:99 – 105.
Harrison, F. W., Warner, B. G. 1986. Fossil fresh-water sponges
(Porifera: Spongillidae) from Western Canada: An overlooked
group of Quaternery paleoecological indicators. Transactions of
the American Microscopical Society 105:110 – 120.
Harrison, F. W., Harrison, M. B. 1977. The taxonomic and ecological status of the environmentally restricted spongillid species of
North America. II. Anheteromeyenia biceps (Lindenschmidt,
1950). Hydrobiologia 62:107 – 111.
Harrison, F. W., Harrison, M. B. 1979. The taxonomic and ecological status of the environmentally restricted spongillid species of
North America. IV. Spongilla heterosclerifera Smith 1918. Hydrobiologia 62:99 – 105.
Harrison, F. W., Johnston, L., Stansell, D. B., McAndrew, W. 1977.
The taxonomic and ecological status of the environmentally
restricted spongillid species of North America. I. Spongilla
sponginosa Penney 1957. Hydrobiologia 53:199 – 202.
Harsha, R. E., Francis, J. C., Poirrier, M. A. 1983. Water temperature: A factor in the seasonality of two freshwater sponge
species, Ephydatia fluviatilis and Spongilla alba. Hydrobiologia
102:145 – 150.
Hartman, W. D., Reiswig, H. M. l973. The individuality of sponges,
in: Boardmann, R. S., Cheetham, A. H., Oliver, W. A. Eds., Animal colonies. Dowden, Hutchinson and Ross, Stroudsburg, PA,
pp. 567 – 584.
Ilan, M., Dembo, M. G., Gasith, A. 1996. Gemmules of sponges
from a warm lake. Freshwater Biology 35:165 – 172.
Jackson, J. B. C. 1977. Competition on marine hard substrata: the
adaptive significance of solitary and colonial strategies. American Naturalist 111:743 – 767.
Jackson, J. B. C., Buss, L. W. 1975. Allelopathy and spatial competition among coral reef invertebrates. Proceedings of the National
Academy of Science, USA. 72:5160 – 5163.
Jewell, M. E. 1935. An ecological study of the fresh-water sponges of
northern Wisconsin. Ecological Monographs 5:461 – 504.
Jewell, M. E. 1939. An ecological study of the fresh-water sponges of
northern Wisconsin, II. The influence of calcium. Ecology
20:11 – 28.
Jones, M. L., Rützler, K. 1975. Invertebrates of the Upper Chamber,
Gaún Locks, Panama Canal, with emphasis on Trochospongilla
leidii (Porifera). Marine Biology 33:57 – 66.
Kratz, T. K., Frost, T.M., Elias, J. E., Cook, R. B. 1991. Reconstruction of a regional, 12,000-year silica decline in lakes using fossil
sponge spicules. Limnology and Oceanography 36:1244 – 1249.
Langenbruch, P-F., Weissenfels, N. 1987. Canal systems and
choanocyte chambers in freshwater sponges (Porifera, Spongillidae). Zoomorphology 107:11 – 16.
Lauer, T. E., Spacie, A. 1996. New records of freshwater sponges
(Porifera) for southern Lake Michigan. Journal of Great Lakes
Research 22:77 – 82.
Mann, K. H., Britton, R. H., Kowalczewski, A., Lack, T. J., Mathews, C. P., McDonald, I. 1972. Productivity and energy flow at
all trophic levels in the River Thames, England, in: Kajak, Z.,
Hillbricht-IlKowska, A. Eds., Productivity problems of freshwaters. PWN, Polish Scientific Publishers, Warsaw, Poland,
pp. 579 – 596
132
Thomas M. Frost, Henry M. Reiswig, and Anthony Ricciardi
McAuley, D. G., Longcore, J. R. 1988. Foods of juvenile ring-necked
ducks: relationships to wetland pH. Journal of Wildlife Management 52:177 – 185.
Mergner, H. 1966. Zum Nachweis der Artspezifität des Induktionsstoffes bei Oscularrohrneubildungen von Spongilliden. Verhandlunger der Deutschen Zoologischen Gesselschaft in Göttingen 30:522 – 564.
Muscatine, L., Porteri, J. W. 1977. Reef corals: mutualistic symbioses
adapted to nutrient-poor environments. Bioscience 27:454 – 460.
Neidhoefer, J. R. 1940. The fresh-water sponges of Wisconsin. Transactions of the Wisconsin Academy of Sciences, Arts, and Letters
32:177 – 197.
Niegel, J. E., Avise, J. C. 1983. Histocompatability bioassays of population structure in marine sponges. Journal of Heredity
74:134 – 140.
Økland, K. A., Økland, J. 1989. The amphiatlantic freshwater sponge
Anheteromeyenia ryderi (Porifera: Spongillidae): taxonomicgeographic implications of records from Norway. Hydrobiologia
171:177 – 188.
Paduano, G. M., Fell., P. E. 1997. Spatial and temporal distribution
of freshwater sponges in Connecticut lakes based upon analysis
of siliceous spicules in dated sediment cores. Hydrobiologia
350:105 – 121.
Palumbi, S. R. 1984. Tactics of acclimation: morphological changes
of sponges in an unpredictable environment. Science
225:1478 – 1480.
Penney, J. T. 1960. Distribution and bibliography (1892 – 1957) of
the freshwater sponges. University of South Carolina Publications, Series 3, 3:1 – 97.
Penney, J. T., Racek, A. A. 1968. Comprehensive revision of a worldwide collection of freshwater sponges (Porifera: Spongillidae).
United States National Museum Bulletin 272.
Pennings, S. C., Pablo, S. R., Paul, V. J. Duffy, J. E. 1994. Effects of
sponge secondary metabolites in different diets on feeding by
three groups of consumers. Journal of Experimental Marine Biology and Ecology 180:137 – 149.
Pile, A. J., Patterson, M. R., Savarese, M., Chernykh, V. I., Fialkov, V.
A. 1997. Trophic effects of sponge feeding within Lake Baikal’s
littoral zone. 2. Sponge abundance, diet, feeding efficiency, and
carbon flux. Limnology and Oceanography 42:178 – 184.
Poirrier, M. A. 1972. Additional records of Texas freshwater sponges
(Spongillidae) with the first record of Radiospongilla cerebellata
(Bowerbank, 1863) from the Western Hemisphere. The Southwestern Naturalist 16:434 – 435.
Poirrier, M. A. 1974. Ecomorphic variation in gemmoscleres of Ephydatia fluviatilis Linneaus (Porifera: Spongillidae) with comments
upon its systematics and ecology. Hydrobiologia 44:337 – 347.
Poirrier, M. A. 1976. A taxonomic study of the Spongilla alba, S.
cenota, S. wagneri species group (Porifera: Spongillidae) with
ecological observations of S. Alba, in: Harrison, F. W., Cowden,
R. R. Eds., Aspects of sponge biology. Academic Press, New
York, NY, pp. 203 – 214.
Poirrier, M. A. 1977. Systematic and ecological studies of Anheteromeyenia ryderi (Porifera: Spongillidae) in Louisiana.
Transactions of the American Microscopical Society 96:62 – 67.
Poirrier, M. A. 1978. Corvospongilla becki N. Sp., A fresh-water
sponge from Louisiana. Transactions of the American Microscopical Society 97:240 – 243.
Poirrier, M. A., Francis, J. C., Labiche, R. A. 1981. A continuousflow system for growing fresh-water sponges in the laboratory.
Hydrobiologia 79:225 – 259.
Poirrier, M. A., Martin, P. S., Baerwald, R. J. 1987. Comparative
morphology of microsclere structure in Spongilla alba, S.
cenota, and S. lacustris (Porifera: Spongillidae). Transactions of
the American Microscopical Society 106:302 – 310.
Proctor, H. C., Pritchard, G. 1990. Variability in the life history of
Unionicola crassipes, a sponge-associated water mite (Acari:
Unionicolidae) Canadian Journal of Zoology 68:1227 – 1232.
Pronzato, R., Manconi, R. 1995. Long-term dynamics of a freshwater sponge population. Freshwater Biology 33:485 – 495.
Racek, A. A., Harrison, F. W. 1975. The systematic and phylogenetic
position of Palaelospongilla chubutensis (Porifera: Spongillidae). Proceedings of the Linnean Society of New South Wales
99:157 – 165.
Randall, J. E., Hartman, W. D. 1968. Sponge-feeding fishes of the
West Indies. Marine Biology 1:216 – 225.
Rasmont, R. 1962. The physiology of gemmulation in fresh-water
sponge. Sympsoium of the Society for the Study of Development
and Growth 20:3 – 25.
Reisser, W. 1984. The taxonomy of green algae endosymbiotic in ciliates and a sponge. British Phycological Journal 19:309 – 318.
Reiswig, H. M. 1970. Porifera: sudden sperm release by tropical
Demospongiae. Science 170: 538 – 539.
Reiswig, H. M., Browman, H. I. 1987. Use of membrane filters for
Microscopid preparations of sponge spicules. Transactions of
the American Microscopical Society 106:10 – 20.
Reiswig, H. M., Ricciardi, A. 1993. Resolution of the taxonomic
status of the freshwater sponges Eunapius mackayi, E. Ingloviformis, and Spongila johanseni (Porifera: Spongillidae).
Transactions of the American Microscopical Society 112:
262 – 279.
Reiswig, H. M., Miller, T. L. 1998. Freshwater sponge gemmules survive months of anoxia. Invertebrate Biology 117:1 – 8.
Resh, V. H. 1976. Life cycles of invertebrate predators of freshwater sponge, in: Harrison, F. W., Cowden, R. R. Eds., Aspects of
sponge biology, Academic Press, New York, NY, pp.
299 – 314.
Resh, V. H., Morse, J. C., Wallace, I. D. 1976. The evolution of the
sponge feeding habit in the caddisfly genusCeraclea (Trichoptera: Leptoceridae). Annals of the Entomological Society of
America 69:937 – 941.
Ricciardi, A., Reiswig, H. M. 1992. Spongilla heterosclerifia Smith,
1918 is an interspecific freshwater sponge mixture (Porifera,
Spongillidae). Canadian Journal of Zoology 70:352–354.
Ricciardi, A., Reiswig, H. M. 1993. Freshwater sponges (Porifera,
Spongillidae) of eastern Canada: taxonomy, distribution, and
ecology. Canadian Journal of Zoology 71:665 – 682.
Ricciardi, A., Snyder, F. L., Kelch, D. O., Reiswig, H. M. 1995.
Lethal and sublethal effects of sponge overgrowth on introduced dreissenid mussels in the Great Lakes — St. Lawrence
River System. Canadian Journal of Fisheries and Aquatic Sciences 52:2695 – 2703.
Richelle, E., Degoudenne, Y., Dejonghe L., Van de Vyver, G. 1995.
Experimental and field studies on the effect of selected heavy
metals on three freshwater songe species: Ephydatia fluviatilis,
Ephydatia mueleri and Spongilla lacustris. Archv für Hydrobiologie 135:209 – 231.
Rioja, E. 1953. Datos historicos acerca de las esponjas de agua dulce
de Mexico. Revista de La Sociedad Mexicana de Historia Natural 14:51 – 57.
Saller, U. 1990. Formation and construction of asexual buds of the
freshwater sponge Radiospongilla cerebellata (Porifera,
Spongillidae). Zoomorphology 109:295 – 301.
Sand-Jensen, K., Pedersen, M. F. 1994. Photosynthesis by symbiotic
algae in the freshwater sponge, Spongilla lacustris. Limnology
and Oceanography 39:551 – 561.
Santelices, B. 1999. How many kinds of individual are there. Trends
in Ecology and Evolution 14:152 – 155.
Schmidt, I. 1970. Phagocytose et pinocytose chez les Spongillidae.
Zeitschrift für vergleichende Physiologie 66:398 – 420.
4. Porifera
Schwandes, L. P., Collins, M. E. 1994. Distribution and significance of freshwater sponge spicules in selected Florida Soils.
Transactions of the American Microscopical Society
113:242 – 257.
Simpson, T. L. 1963. The biology of the marine sponge Microciona prolifera (Ellis and Solander): I. A study of cellular function and differentiation. The Journal of Experimental Zoology 154: 135–151.
Simpson, T. L. l973. Coloniality among the Porifera, in: Boardmann,
R. S., Cheetham, A. H., Oliver, W. A. Eds., Animal colonies.
Dowden, Hutchinson and Ross, Stroudsburg, PA, pp. 549 – 565.
Simpson, T. L. 1984. The cell biology of sponges. Springer-Verlag,
New York, NY.
Simpson, T. L., Fell, P. E. 1974. Dormancy among the Porifera: gemmule
formation and hatching in freshwater and marine sponges. Transactions of the American Microscopy Society 93:544–577.
Simpson, T. L., Gilbert, J. J. 1973. Gemmulation, gemmule hatching,
and sexual reproduction in fresh-water sponges: I. The life cycle
of Spongilla lacustris and Tubella pennsylvanica. Transactions
of the American Microscopy Society 92:422 – 433.
Smith, D. G. 1994. First report of a fresh-water sponge (Porifera:
Spongillidae) from the West Indies. Journal of Natural History
28:981 – 986.
Smith, L. C., Hildemann, W. H. 1984. Alloimmune memory is absent
in Hymeniacidon sinapium, a marine sponge. The Journal of
Immunology 133:2351 – 2355.
Sowka, P. A. 1997. Occurrence of two species of freshwater sponges
(Dosilia radiospiculata and Ephydatia muelleri) in Arizona.
Southwestern Naturalist 44:211–212.
Stoddart, J. A., Ayre, D. J., Willis, G., Heyward, A. J. 1985. Selfrecognition in sponges and corals? Evolution 39:461 – 463.
Ungemach, L. F., Souza, K., Fell, P. E., Loomis, S. H. 1997. Possession and loss of cold tolerance by sponge gemmules: a comparative study. Invertebrate Biology 116:1 – 5.
Vacelet, J., Boury-Esnaault, N. 1995. Carnivorous sponges. Nature
373:333 – 335.
Vacelet, J., Boury-Esnaault, N., Fiala-Medioni, A., Fisher, C. R.
1995. A methanotrophic carnivorous sponge. Nature 377:296.
van Weel, P. B. 1949. On the physiology of the tropical fresh-water
sponge Spongilla proliferens Annand. I. Ingestion, digestion,
and excretion. Physiolgia Comparata et Oecologia 1:110 – 128.
Van de Vyver, G. 1970. La non confluence intraspécifique chez les
spongiaires et la notion d’individu. Extrait des Annales d’Embryologie et de Morphogenèse 3:251 – 262.
Van de Vyver, G. 1979. Cellular mechanisms of recognitions and
rejection among sponges. in: Levi, C., Boury-Esnault, N. Eds.,
Biologie des Spongiaires. Colloques Internationaux du Centre
National De La Recherche Scientifique, No. 291, Paris, France,
pp. 195 – 204.
Van de Vyver, G., Willenz, Ph. 1975. An experimental study of the
133
life cycle of the fresh-water sponge Ephydatia fluviatilis in its
natural surroundings. Wihelm Roux’ Archiv 177:41 – 52.
Vogel, S. 1974. Current induced flow through the sponge, Halichondria. Biological Bulletin (Woods Hole, Massachusetts) 147:
443 – 456.
Volkmer-Ribeiro, C. 1981. Porifera. in: Hurlbert, S. H., Rodriquesz,
G., Santos, N. D. Eds., Aquatic biota of tropical South America,
Part 2: Anarthropoda. San Diego State University, San Diego,
CA, pp. 86 – 95.
Volkmer-Ribeiro, C. 1986. Evolutionary study of the freshwater
sponge genus Metania GRAY, 1867: III. Metaniidae, new family. Amazoniana 9:493 – 509.
Volkmer-Ribeiro, C., Rosa-Barbosa, R. De. 1979. Neotropical freshwater sponges of the Family Potamolepidae Brien, 1967. in:
Levi, C., Boury-Esnault, N. Eds., Biologie des Spongiaires. Colloques Internationaux du Centre National De La Recherche Scientifique, No. 291, Paris, France., pp. 503 – 511.
Volkmer-Ribeiro, C., Grosser, K. M. 1981. Gut contents of Leporinus obtusidens “sensu” Von Ihering (Pisces, Characoidei) used
in a survey for freshwater sponges. Revista Brasileira de Biologia 41:175 – 183.
Weissenfels, N. 1975. Bau und Funktion des Süsswasserschwamms
Ephydatia fluviatilis L. (Porifera). II. Anmerkungen zum Körperbau. Zeitschrift für Morphologie der Tiere 81:241 – 256.
Weissenfels, N. 1976. Bau und Funktion des Süsswasserschwamms
Ephydatia fluviatilis L. (Porifera). III. Nahrungsaufnahme, Verdauung und Defäkation. Zoomorphologie 85:73 – 88.
Weissenfels, N. 1980. Bau und Funktion des Süsswasserschwamms
Ephydatia fluviatilis L. (Porifera). VII. Die Porocyten. Zoomorphologie 95:27 – 40.
Weissenfels, N. 1981. Bau und Funktion des Süsswasserschwamms
Ephydatia fluviatilis L. (Porifera). VIII. Die Entstehung und Entwicklung der Kragengeissselkammern und ihre Verbindung mit
dem ausführenden Kanalsystem. Zoomorphology 98:35 – 45.
Weissenfels, N. 1982. Bau und Funktion des Süsswasserschwamms
Ephydatia fluviatilis L. (Porifera). IX. Rasterelektronmikrosckpische Histologie und Cytologie. Zoomorphology 100:75 – 87.
Weissenfels, N. 1990. Condensation rhythm of fresh-water sponges
(Spongillidae, Proifera). European Journal of Cell Biology
53:373 – 383.
Weissenfels, N. 1992. The filtrations apparatus for food collection in
freshwater sponges (Porifera, Spongilidae). Zoomorphology
112:51 – 55.
Willenz, P. 1980. Kinetic and morphological aspects of particle ingestion by the freshwater sponge Ephydatia fluviatilis L., in: Smith,
D. C., Tiffon, Y. Eds., Nutrition in lower Metazoa, Pergammon
Press, Oxford, England, pp. 163 – 187.
Williamson, C.W. 1979. Crayfish predation on freshwater sponges.
American Midland Naturalist 101:245 – 246.
This Page Intentionally Left Blank
5
CNIDARIA
Lawrence B. Slobodkin
Patricia E. Bossert
Department of Ecology and Evolution
State University of New York
Stony Brook, New York 11794
Science Department
Northport High School
Northport, New York 11768
I. Introduction
II. General Biology of Cnidaria
A. Body Plan
B. Nematocysts
C. Feeding
D. Reproduction and Metamorphoses
E. Ecological Interactions
III. Descriptive Ecology of Freshwater Cnidaria
A. Hydra
B. Craspedacusta
C. Calposoma
D. Cordylophora
E. Polypodium Hydriforme
IV. Collection and Maintenance of Freshwater
Cnidaria
A. Collecting Techniques
B. Maintenance Procedures
V. Classification of Freshwater Cnidaria
A. Species Groups of Hydras
B. Taxonomic Key to Genera of
Freshwater Cnidaria
Literature Cited and Further Readings
I. INTRODUCTION
Medusae, anemones, corals, and other polyps
compose the ancient and remarkably successful phylum, Cnidaria. They occur as fossils in the lithographic stone of the Mid-Cambrian Burgess shale,
and have not changed very dramatically since then.
Fossilized cnidarian embryos are reported from the
lower Cambrian (Kouchinsky et al., 1999). The phylum name, Cnidaria, is derived from the Greek term
for “nail”, based on their possession of nematocysts,
which look like rods attached to a round capsule. The
other name for the phylum is Coelenterata, a term
alluding to their saclike internal space, the coelenteron. A general overview of the phylum and survey
of older literature is provided by Hyman (1940) and
the papers in various conference proceedings (e.g.,
Mackie, 1976).
The freshwater representatives of Cnidaria are
small animals belonging to the class Hydrozoa, with
relatively few species and somewhat monotonous
anatomy. They consist of the following taxa:
1. The common and familiar hydras, a group of
secondarily simple, solitary polyps;
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
2. The sporadically common Craspedacusta and
Limnocodium, jellyfish with minute, polypoid
larvae (Boulenger and Flower, 1928; Fuhrman,
1984).
3. Calposoma, a tiny, colonial polyp, which is so
small and inconspicuous that it is probably
more common than it appears to be (Rahat and
Campbell, 1974; Fuhrman, 1984).
4. Polypodium, which spends part of its life cycle
as a parasite in the eggs of sturgeon and part as
an ambulatory, predaceous polyp (Lipin, 1926;
Raikova, 1973, 1980); this species has been
described primarily from eastern European
rivers, but should also be found in North
American sturgeon.
5. Various estuarine coelenterates, which may
occasionally occur in relatively fresh water;
colonial, sessile animals of the genus Cordylophora will serve as examples of these
(Hubschman and Kishler, 1972; Roos, 1979).
Hydra and Cordylophora belong to the order
Hydroida of the class Hydrozoa. Craspedacusta, Limnocnida, and Calposoma are classified as members of
the order Limnomedusae. The parasitic Polypodium
135
136
Lawrence B. Slobodkin and Patricia E. Bossert
has been assigned to the order Trachylina of the
same class (cf. Hyman, 1940). The fact that the other
three orders of the class Hydrozoa (Actinulida,
Siphonophora, and Hydrocorallina) and the other
three classes of coelenterates (Scyphozoa, Cubozoa,
and Anthozoa) are not represented outside of the sea is
curious. There is not even any serious speculation as to
why freshwater invasion by coelenterates has been so
severely limited. There is no special osmoregulatory
organ in the phylum, but this is not an explanation
since its absence has not stopped the successful invaders. Because of their small size, soft bodies, and
often sessile habits, freshwater Cnidaria are either not
collected or not well preserved in most routine collecting procedures. They are, however, widely distributed
and can be found in most ponds and streams when a
specific search is made. When they are abundant, they
can be major predators of small invertebrates and even
of tiny fish. In turn, Hydra is fed upon by flatworms,
and crayfish eat Craspedacusta. Probably, other animals prey on freshwater Cnidaria, but this has not been
carefully studied.
II. GENERAL BIOLOGY OF CNIDARIA
A. Body Plan
All cnidarians share a simple body plan of a central
cavity surrounded by two cellular layers (Fig. 1). The
endoderm lines the interior cavity, the coelenteron.
Between the endoderm and the ectoderm is an intermediate, noncellular mesoglea. Nematocysts aid in feeding
and in repelling predators, and are present in all freshwater and marine cnidarians.
FIGURE 1 Cross section through digestive cavity (coelenteron) of
generalized cnidarian.
The feeding aperture or mouth leads into the coelenteron, which functions as a gut. By convention, the
end of the animal with the mouth is termed “oral”, the
opposite end “aboral.” The oral aperture serves at different times as mouth and anus. When the mouth is
closed, the pressure of fluid in the coelenteron can
stiffen the body, even in the complete absence of any
hard tissue. The coelenteron, therefore, functions also
as a hydrostatic skeleton.
The mesoglea varies enormously in thickness
among different members of the group. At its most
meager, as in Hydra, it is not more than 200 m thick,
containing only wandering cells, nonliving fibrous
components, and fibers from neuromuscular cells. It is
more fully developed in medusae, such as Craspedacusta, and may contain a great deal of collagenous or
gelatinous material. The ectodermal body wall has
neural and contractile properties and is also the location for ripe nematocysts.
The presence of definite cell layers with differentiated functions distinguishes these animals as true metazoa. The absence of mesoderm implies that they do not
have organs like higher metazoa; therefore, such terms
as “tentacles” and “gut” are used in a functional sense.
The basic body plan can be manifested as either a
polyp or a medusa (Fig. 2). Polyps are typically elongated along the oral – aboral axis. Medusae are approximately bell-shaped and usually have their greatest
body dimension perpendicular to the oral – aboral axis.
The coelenteron of a polyp is usually deeper than its
body is wide, while a medusa is usually wider than
its coelenteron is deep. Also, medusae generally have
relatively thicker mesoglea. In some species, different
generations, and, in some cases, the same individual organism at different stages of its development, can adopt
the form of either a polyp or a medusa.
Around the feeding aperture of polyps or the edge
of the bell of medusae is typically a ring of tentacles.
Tentacles are extensions of the two cellular layers into
more or less elongated projections. The coelenteron
FIGURE 2
(B) medusa.
Basic body plan of Cnidaria showing (A) polyp and
5. Cnidaria
may or may not extend into the tentacles. Tentacular
ectoderm is especially rich in nematocysts, which may
be arranged in rosette or ring-shaped batteries. Tentacles are used in food capture, defense, and in some
cases, locomotion.
B. Nematocysts
The phylum is characterized by the presence of
cnidoblasts, ectodermal cells that produce the cell
products called cnida or nematocysts. Because there are
many kinds of elaborately spined nematocysts, these
structures are valuable characters for the classification
of coelenterates, particularly in such morphologically
monotonous groups as Hydra.
Nematocysts consist of a capsule containing a
threadlike tube (Fig. 3A). Near the base of the tube is a
projection reminiscent of a trigger. Remnants of the living cnidoblast are absent from mature nematocysts.
Many nematocysts come to lie on the tentacles, often
after having been moved through the mesoglea from
some other region.
After firing, a nematocyst consists of a long thread
with a capsule at its base (Fig. 3B). Some nematocysts
FIGURE 3
137
(the stenoteles or penetrants) are open at the thread tip,
giving the appearance of a hypodermic syringe with a
long needle. Stenoteles may have a complex of thornlike structures around the base of the thread. Stenoteles
eject a neurotoxin, which partially paralyzes the prey.
Desmonemes (or volvonts), another form of nematocyst, seem to lack poison, but rather eject sticky
threads that wind about the spines and hairs on the
body of the prey, interfering with movement. The capsules of some volvonts remain fixed to the tentacles
after firing so that their exploded threads fasten prey to
the tentacles as if by many tiny ropes or grappling
hooks.
At least 17 morphologically distinct forms of nematocysts are present in the phylum (Fig. 4). The penetrants and volvonts of Hydra come in several forms,
classified in terms of capsule size, spination of the
threads, and shape and distribution of the basal spines.
A single animal may have five or more types of nematocysts. Nematocysts of basically the same type may differ
among species in how the thread is coiled inside the
capsule prior to eversion. Some may appear like a coiled
spring, with gyres at right angles to the longest dimension of the capsule, while others are coiled parallel to
Discharged nematocyst from Hydra.
138
Lawrence B. Slobodkin and Patricia E. Bossert
making them of interest as examples of complex cell
differentiation. Also, the mechanism by which the thin
thread of the nematocyst everts from its coiled state
within the capsule (like an enormously elongated, inverted finger of a glove suddenly turning itself inside
out) is a difficult problem in fluid pressures. The poison
that is secreted by some nematocysts through the open
tip of the thread is of medical interest. Furthermore,
the enormous diversity of shapes, sizes, and spination
of nematocysts within single organisms and among the
different coelenterates, poses a problem in cell differentiation and genetics. Readers interested in more details
on nematocysts should consult the large review volume
by Hessinger and Lenhoff (1988), which is too lengthy
to summarize here.
C. Feeding
FIGURE 4
Some of the types of nematocysts present in Cnidaria
(redrawn from Hyman, 1940); (A) rhopaloneme (only in
Siphonophora); (B) spirocyst; (C) same as B, unraveling (not discharged); (D) desmoneme [(H) and (J) same as D but not discharged];
(E) atrichous hydrorhiza [(L) same as E, but discharged]; (F) holotrichous isorhiza [(K) same as F but discharged]; (G) stenotele inside
its cnidoblast [(M) same as G but discharged]; (N) microbasic
amastigophore; (O) homotrichous microbasic eurytele; (P) heterotrichous microbasic eurytele; (Q) macrobasic mastigophore; (R) teleotrichous macrobasic eurytele; (S) heterotrichous anisorhiza; and (T) microbasic mastigophore. 1, Capsule; 2, tube; 3, butt; 4, cnidoblast; 5,
its nucleus; 6, lid; and 7, stylet.
the long dimension of the capsule. Nematocyst structure was initially considered a central taxonomic character; but recent evidence indicates that details of shape
and coiling, and also the relative abundance of nematocysts of different types, are somewhat variable even
within clones of Hydra (Campbell, 1987).
The microanatomy and function of nematocysts
is a rich research area. The enormous interest about
nematocysts and their production arises in part from
the following observations. On the level of electron
microscopy, they are extremely elaborate structures
After a prey has been stung and encumbered by the
nematocysts, the tentacles move the victim to the
mouth, which opens to admit it into the coelenteron.
Often the prey is still alive and active, but this ceases as
soon as the gastric cells lining the coelenteron secrete
digestive juices.
Since there is no anus, food cannot be passed along
the gut while digestion continues (as in higher metazoa).
Instead, feeding stops until the digestion process is completed and the indigestible remnants have been regurgitated. In hydra, ingested food decomposes within an
hour into a slurry of particles. The role of food vacuoles
is reminiscent of the feeding process in protozoa and
sponges (Barnes, 1966). The free borders of the digestive
cells ingest particles by pinocytosis. Individual food particles are enclosed in vacuoles that are moved through
endodermal cells. Eventually, their indigestible residues
are ejected by the endodermal cells and returned to the
coelenteron to leave ultimately through the mouth.
D. Reproduction and Metamorphoses
In the coelenterates, many kinds of reproduction
exist (Hyman, 1940). Like all metazoa, they can reproduce sexually. The fertilized eggs may produce larvae
differing anatomically and ecologically from the adult
sexually reproducing stage. In marine coelenterates,
there may be an elaborate succession of larval stages,
some of which may form colonies or reproduce
vegetatively by budding or fragmentation. In any particular species, one or more of these stages may be
missing. There are medusae that produce eggs that
go through various larval stages to produce new
medusae (Fig. 5A). Some polyps produce gonads
(Fig. 5A), and others bud off medusae that either swim
away to become sexual (Fig. 5C) or remain attached to
5. Cnidaria
139
FIGURE 5
(A) Sexual medusa produces larva; larva forms new medusa; (B) sexual polyp generates new
polyp; and (C) polyp produces medusa which becomes sexual and forms new polyp [redrawn from Barnes,
1966.]
their parents and become sexual without ever feeding
independently.
Polyps or medusae may produce new individuals
that may or may not resemble their “parent.” A new
individual may separate from its parent; but in many
coelenterates, the asexually produced individuals stay
attached and form a colony. The development of
colonies is absent in hydra, but occurs among all other
freshwater Cnidaria.
In the anatomically simplest coelenterate colonies,
such as the colonial microhydra, “larvae” of the freshwater Craspedacusta, all attached individuals are essentially similar in both form and function (Payne, 1924).
Each has its own mouth and coelenteron and each is
capable of budding new polyps. Microhydra larvae can
produce medusae by budding. Colonies of many coelenterates are much more elaborate.
In a common modification of this process, certain
members of the colony become “gonozooids” that neither feed nor have tentacles; instead, they consist of a
stalk rising from the common stolon that buds off
medusae. These medusae, in turn, either form gonads
while still attached to their colony or leave the colony
and then produce gonads.
The best known freshwater coelenterates, the hydra, have lost the medusa stage entirely. In hydra,
polyps may produce new polyps by asexual budding or
may temporarily switch to sexual reproduction using
140
Lawrence B. Slobodkin and Patricia E. Bossert
gonads. Fertilized eggs derived directly from polyps
may then hatch to produce new polyps. There are several marine examples, but only one freshwater example
of medusae budding new medusae. This is in a species
presumed endemic to Lake Tanganyika, in the great
African rift (Thiel, 1973).
E. Ecological Interactions
Although some species may gain part of their nourishment from intracellular algal symbionts, all cnidarians are carnivores. Their prey consists primarily of
small coelomate animals, organisms that evolved long
after the appearance of coelenterates. They generally
do not feed on protozoa, nematodes, or sponges, nor
will these animals trigger the nematocysts. This raises a
curious question, for which there is no evident answer:
“What did the first coelenterates eat?”
Hydra are extremely effective predators. Prey includes crustaceans, worms, and fish and insect larvae.
They have been seriously considered as a biological control organism for mosquitoes. Hydra and Craspedacusta
can both sting fishes very badly. Fish that are too big to
swallow often cannot survive being stung. For this reason, hydra are sometimes serious pests of fish hatcheries.
Nematocysts are sufficiently unpleasant that relatively few predators attack coelenterates. Turtles, fish,
crabs, worms, echinoderms, and flatworms are among
the predators on marine coelenterates. Predation on the
freshwater coelenterates is not well studied. Crayfish
eat Craspedacusta (Dodson and Cooper, 1983). Flatworms are reported to eat Hydra (Hyman, 1940;
Kanaev, 1969), but we have seen small flatworms withdraw from contact with hydras.
Some marine coelomates are not affected by nematocysts and are commensal with coelenterates. On coral
reefs, clown fish live among tentacles of large
anemones and other fish occur only in close association
with the Portuguese man-of-war. In freshwater, Anchistropus, a chydorid cladoceran, has been observed
clinging to, and apparently feeding on, the body wall
of hydras (Griffing, 1965; Slobodkin, personal observation). The amoeba Hydramoeba hydroxena feeds on
hydra (Stiven, 1976), and the hypotrichous ciliate
Kerona lives on the surface of hydras (Hyman, 1940).
Polypodium hydriforme is parasitic on a fish for part
of its life (Raikova, 1980).
III. DESCRIPTIVE ECOLOGY
OF FRESHWATER CNIDARIA
Their anatomical simplicity and the ease with
which hydra can be cultured in the laboratory make
freshwater Cnidaria very important as experimental
material in cellular and developmental biology, as well
as in neurobiology, biochemistry (Ciernichiari et al.,
1969), and genetics (Sabine et al., 1987; Schummer
et al., 1992). Hydra also lends itself to studies in cell
growth, morphogenesis, microanatomy, and symbiosis
(Ciernichiari et al., 1969; Dunn, 1986, 1987, 1988;
Bossert and Dunn, 1996). In the present chapter, we
will focus on cnidarian ecology and natural history,
which have been less well studied.
A. Hydra
1. General Biology of Hydra
Hydra are small polyps from 1 to 20 mm in body
length. The body is crowned by up to 10 or 12 tentacles. Usually the tentacles are of approximately the
same length as the body, but may be somewhat shorter,
particularly in the green hydra, and can exceed 20 cm
in length in hungry brown hydra in quiet water. There
are no medusae. Reproduction is by budding or by gametes produced directly from ectoderm. The fertilized
eggs may enter a resting stage. When development
proceeds, the egg immediately develops into a polyp
(see Fig. 5).
Hydra are found attached to almost any reasonably hard surface. Slight bacterial films may make surfaces more attractive, but heavy growth of microalgae
may be avoided. Water lily stems, charophytes, dead
leaves, sticks, and stones are favored substrates. They
may appear as single animals or as dense furlike aggregations. There are reports of fishing nets becoming
completely covered with brown hydra, resulting in
rashes on the hands and arms of the fishermen (Batha,
1974).
If the wait for food is longer than approximately
12 h, the hydra begin to change their locations on the
substrate (Ritte, 1969; Lenhoff and Lenhoff, 1986).
They have two different methods of movement. Smallscale movements on a substrate may occur by attaching
the stretched tentacles to the substrate or, in shallow water, to the surface film, releasing the pedal attachment
and contracting the tentacles and reattaching. Sufficiently crowded or hungry hydras float off their substrate (Lomnicki and Slobodkin, 1966). They may appear in the plankton or be found floating upside down
with the pedal disk in the water surface film and their
tentacles trailing.
Batha (1974) extensively documented the existence
of planktonic hydra in Lake Michigan through use of
divers and of suitable attachment surfaces suspended in
midwater. In addition, the reports of fishnets covered
by hydra and observations by Griffing (1965) of
5. Cnidaria
sudden relocations of hydras within a single pond suggest that hydras are probably much more important
components of lake plankton than has been generally
realized.
Floating animals will sink and settle either from
wave and current action or from having just fed. This
behavior keeps hydra in areas of abundant food. It is
also evolutionarily important as a dispersal mechanism.
From the perspective of a naturalist, it has the effect of
making it relatively easy to collect hydra at the outflow
of lakes and ponds.
They are not tolerant of heavy metals, but they can
thrive in even highly eutrophic water. They can survive
at temperatures from near freezing to 25°C.
Several Hydra species may coexist in a pond; often
a small green hydra and, at least, one large brown
species will co-occur. Also, strains of Hydra may replace each other seasonally. Bossert (1988) showed that
green hydra collected several months apart from the
same small pond had very different size and growth
characteristics when maintained under very similar
conditions in the laboratory.
Despite many years of collection and observation,
we have never found a species of green hydra that was
consistently larger than any strain of asymbiotic brown
hydra, nor have we found any contradictory account in
the scientific literature. If it occurs, it certainly seems
rare. Some large brown hydra can be caused to become
green in the laboratory (Rahat and Sugiyama, 1993),
but it is not clear that this is significant in nature.
Hydra occur in freshwater from the Amazon to
Alaska and from Siberia to Africa at depths from shallow water to 60 m or more. This cosmopolitan distribution may be a result of the portability of the thecate
eggs; or perhaps, as has been suggested by Campbell
(1987), it might be due to the four species groups (see
Section V) having differentiated before the primeval
continental masses separated in the Mesozoic era. A recent study of the hydras of Madagascar (Campbell,
1999), which has been an island for more than 100
million years, shows that they differ very little from
mainland hydra, indicating that either the rate of
evolution of the genus Hydra is very slow or that they
are surprisingly well dispersed. One species group
(oligactis) is missing from Madagascar. Since there is
no obvious reason why hydra of this group should be
less mobile than the others, their absence strengthens
the interpretation of slow evolution. This implies that
hydras are evolutionarily very old.
2. Reproduction and Mortality
Asexual budding is the primary reproductive mechanism of hydra during periods of population increase.
Under optimal conditions of food supply, temperature,
141
FIGURE 6
Hydra. (A) Male gonad; (B) female gonad; and (C) egg
[redrawn from Campbell, 1987.]
and water quality, each adult hydra polyp can produce
two buds per day and each bud can mature and begin
reproducing in a week or less. Generally, the smaller
strains of hydra bud more rapidly than the larger ones,
at equal feeding rates. Buds are from 12 to 20% of the
size of the mother, varying with the strain.
While Hydra usually reproduce by producing freeliving buds, under deleterious environmental conditions they may develop testes or single egg ovaries and
engage in sexual reproduction. External fertilization by
free-swimming sperm occurs while the egg is attached
to the body wall of the mother. The embryo may enter
a resting period of days or even months before proceeding with direct development into a new polyp.
During sexual reproduction, gonads develop along
the stalk in place of buds (Fig. 6). Mature male gonads
are a mound of tissue with a distinct apical nipple from
which sperm extrude. Mature female gonads consist of
a single large egg cell resting on a cushion of smaller
cells. The fertilized eggs are surrounded by a theca,
which may be smooth or ornamented or may consist of
polyhedral plates. The features of the theca are of taxonomic importance.
Sexuality in hydra seems to occur only under deleterious environmental conditions, although the precise
cues are unknown. Chemical changes due to temperature fluctuations, and, perhaps, sudden nutritional variation can all induce sexuality at certain times (Lenhoff
and Loomis, 1961; Kanaev, 1969). Individual hydra
have been reported as unisexual and some bisexual
individuals have been noted (Hyman, 1940; Kanaev,
1969). If conditions improve, sexual hydra can return
142
Lawrence B. Slobodkin and Patricia E. Bossert
to asexual reproduction. Buds and gonads may occur
concurrently in the same individual.
Most studies of hydra sexuality are made on animals that have been maintained in the laboratory.
Batha (1974) has suggested that sexual reproduction is
very rare in nature, based on the complete absence of
gonads among thousands of brown hydra collected in
Lake Michigan. Although we have found eggs in green
hydra collected from a small pond and other field reports of gonads do exist (Kanaev, 1969), it seems clear
that in hydra the numerically most important reproductive process is budding.
In most plants and animals, population size and
genetic recombination are associated through the
process of sexual reproduction, but sexual reproduction in hydra is not significant to population dynamics.
It seems important for maintaining genetic heterogeneity and permitting escape from temporarily unsuitable
conditions (cf. the chapter on “Escape in Time and
Space” in Slobodkin, 1980).
In a single pond or stream, most of the animals are
likely to belong to a rather small number of vegetative
reproductive lines, being genetically identical except for
occasional mutations. Even those that have emerged
from eggs are likely to be the result of relatively close
inbreeding, since sexuality is usually found in crowded
local populations whose members are descended from
a very small number of clonal lines. There is no direct
evidence on sperm survival in nature, but it seems unlikely that sperm can travel great distances.
Hydra that are very small, either because of youth
or starvation, will not produce buds. If nutrition is limited, animals may be kept indefinitely in a condition of
neither growing nor budding. Both body size and budding rate are proportional to feeding rate up to a point
of food saturation.
The capacity of hydra to reduce its body size is of
ecological interest. Animals with hard skeletons are
committed to a particular body size in the sense that
they cannot shrink below a given point during periods
of food shortage. The inability to reduce size may contribute to death by starvation. By contrast, hydras
shrink in size when starved, but can be restored to full
reproductive size by increasing their food supply. At a
given temperature, hydras will come to a steady-state
body size and budding rate if food supply is sufficient
and constant.
The time for complete size adjustment to either
temperature change or feeding level change on the part
of an individual brown hydra is approximately 3
weeks. Reduction of temperature decreases growth
rate, but increases body size. These size differences
seem to be due to changes in cell number rather than
cell size (Fig. 7) (Hecker and Slobodkin, 1976). Because
FIGURE 7 Relative sizes of hydra raised at different temperatures.
Scale shown is the same size at each temperature [redrawn from
Hecker and Slobodkin, 1976.]
larger animals have greater food requirements, the
smaller green hydra can produce buds when fed one or
two Artemia salina nauplii per day while the larger
brown hydra require 5 – 10 per day before they will
bud.
3. Feeding
The tentacles extend above and lateral to the body.
They are generally motionless except in the presence of
potential food or during locomotion. In healthy animals, the tentacles are cylindrical or tapered, but never
clubbed. The length of the tentacles varies somewhat
with species, but considerably more with environmental circumstances. In quiet water, animals with bodies
no longer than 2 cm have been observed to constrict
their tentacles into thin threads extending at least
20 cm (L. B. Slobodkin, personal observation). This is
unusual however, and tentacles equal to three body
lengths or less are more typical.
Tentacles are quickly retracted if the animal is disturbed or if organisms brush against them. Whole prey
or extracts of their body fluids will initiate active waving movements of the tentacles (Lenhoff, 1983). If a
prey organism brushes against the tentacles, nematocysts will discharge, poisoning the prey and attaching it
to the tentacle. Other tentacles and their nematocysts
then join the attack. In a matter of 1 – 4 min, the prey
will have been pressed against the mouth surface by the
tentacles and will have entered the coelenteron. Meanwhile, the tentacles for subsequent swallowing may
have caught other prey.
The number of prey swallowed in one feeding encounter varies with the size of the hydra, the size of the
5. Cnidaria
prey, and the previous feeding condition of the hydra.
One large Daphnia magna may fill the coelenteron
completely, stretching its walls to give the appearance
of a Daphnia stuffed into a thin expandable sack of
hydra tissue.
During the digestion process, the body may become rounded as the swallowed prey are reduced to a
slurry. After approximately 1 h, material is suddenly
discharged from the coelenteron through the mouth,
the hydra momentarily appearing like a punctured balloon. The columnar shape is then restored, tentacles
regain their virulence, and the animal waits for its next
meal. There is evidence that several days of starvation
will increase the appetite of a brown hydra. Brown hydra can survive more than 40 days without food, and
green hydra can live without food 4 months or longer.
In general, hydra eat small, open-water plankters,
but are less effective at capturing animals that normally
inhabit underwater surfaces. The common cladoceran
genera Simocephalus, Scapholebris, and Chydoris and
at least some ostracods are immune to the activities
of hydra (Schwartz et al., 1983). Hydra can eat very
small fish and insects, but sufficiently large animals
with hard skeletons and strong swimming force can escape after being stung. The long bristles on small midge
larvae have been found to impede predation by hydra
(Hershey and Dodson, 1987).
Apparently, hydra primarily feed on the kinds of
prey that they are least likely to encounter. This suggests that hydras are sufficiently important as natural
predators that only those crustaceans that have evolved
immunity to hydra can coexist closely with them.
4. Immortality and Regeneration
Trembley, in the eighteenth century, had already
demonstrated the capacity of hydra to regenerate perfectly, even after severe mutilations. The regenerative
powers of hydra make them favorite classroom objects.
They can be decapitated, bisected, have their tentacles
amputated, or even be turned inside out and in a matter
of a few days regenerate missing parts or regain their
proper organization. Within a clone, rings of hydra
stalk can be threaded on hairlike quoits and may fuse to
form a single tube (Burnett, 1973). Even without operations, different accidental conditions in the field or laboratory will produce hydra with various mutilations:
more or fewer tentacles, missing heads and so on. All of
these hydras will reorganize themselves neatly if water
chemistry is not deleterious and if they have been reasonably well-nourished before the mutilations occurred.
Since hydras can regenerate so well, are they potentially immortal? Immortality in any organism is impossible to demonstrate during a research program of
finite duration, and therefore the question cannot be
143
unequivocally settled. However, what we generally
mean by immortality in organisms is the absence of any
signs whatsoever of senescence or permanent scars of
the past. Martinez (1997) maintained laboratory cohorts of hydra for more than 3 years and demonstrated
the absence of any symptoms of senescence. Single
polyps have been maintained in our laboratory without
budding for at least a year, their lives terminating only
from human error. There is no clear evidence for senescence of any kind in hydra. If hydra are starved they
will become smaller; but do not die until they are too
small to feed. Newborn hydra are approximately at
this minimum size (Slobodkin et al., 1991).
It seems likely that hydra polyps are potentially immortal. All deaths in hydra can be assigned to such
things as alterations in water quality or food supply,
temperature shocks, excessively severe starvation, or
predation.
5. Symbiosis
While species of Hydra may be difficult to distinguish, there is a very clear distinction between the brilliant green color of some species of Hydra and the
yellow, brown, and gray colors of those that do not
have symbiotic algae. The green hydra are accepted as
a taxonomically distinct group and have been assigned
their own generic status as Chlorohydra, which we use
here interchangeably with “green hydra.” No certainty
exists as to the monophyletic character of this genus.
The green color arises from Chlorella-like cells,
unicellular algae each occupying a vacuole in the
endodermal cells of their hosts. Each endodermal cell
contains 10 – 35 algae-laden vacuoles (Fig. 8). The precise number varies with species, but is nearly constant
within a hydra strain and a set of environmental circumstances (Bossert, 1986). Algae are also found in the
central cells of the tentacles, but in somewhat smaller
numbers. The endoderm of the buds contains algae like
those of the mother.
Ample evidence exists showing that green hydra
gain nourishment from their symbionts. Radioactivetracer experiments have demonstrated that maltose, a
secondary photosynthetic compound, leaks from the
algal symbionts to their hosts (Muscatine, 1965;
Ciernichiari et al., 1969; Lenhoff, 1974). Also, there is
microscopic evidence that algal cells can be attacked by
host lysozymes (Dunn, 1986, 1987, 1988). In competition experiments between brown hydra and green hydra populations, light provides a significant advantage
to the green hydra (Slobodkin, 1964).
The algae can be removed from some strains
of green hydra by use of a variety of techniques, including photosynthetic poisons, prolonged darkness or
extremely strong light, and dilute glycerin solutions
144
Lawrence B. Slobodkin and Patricia E. Bossert
FIGURE 8
An algae-laden endodermal cell from Hydra (viridis) (1000 ).
(Pardy, 1983). Such hydras are referred to as aposymbiotic and are susceptible to reinvasion by algae. It is
possible to develop green patches in brown hydra by
injecting their coelenteron with algae or by feeding
green hydra to brown ones (L. B. Slobodkin, personal
observation), but this coloration fades with time.
If the association with algae is advantageous, why
are big hydras not green? Symbiotic relationships involve a delicate interaction between the two partners,
particularly when one partner lives within the cells of
the other. A primary requirement for stable endosymbiosis is a mechanism providing balanced rates of mitosis of the host and symbiont cells. If there were no control of the algal rate of increase, they would be expected
to kill their host cells by filling them with algae; and if
the hydra cells excessively limited the algae, they would
be eliminated and the hydra would be brown.
While the number of algae per host cell stays constant in all green hydra, maintaining this constancy
seems more difficult in the larger green hydra strains.
Bossert and Dunn (1996) have shown that algal cells
are increasing faster than host cells in green hydra
strains of all sizes, but the disparity between algal and
animal mitotic rates is greatest in the largest strains.
Dunn (1986, 1988) suggested that algal cells are being
actively expelled or digested by all green hydra, but
especially by larger strains.
Size in green and brown hydra is apparently regulated by the relative amounts of several hormones,
some of which activate the formation of buds while
others inhibit budding (Sabine et al., 1987). Bossert
(1988) found that one of the hormones implicated in
producing smaller hydra size also inhibits algal mitosis
within green hydra cells. Apparently, the same hormonal mechanism that produces small size aids in
maintaining the balance between algal cell and hydra
cell increases, while a hormonal balance that permits
larger size makes control of algae more difficult and, in
the largest strains, impossible. More investigation of
this problem is needed.
5. Cnidaria
Green hydras are never found in nature as aposymbionts and seem to have evolved a dependence on algae. However, these algae have probably not evolved a
dependence on hydra. In fact, how the algae benefit
from the association is not at all clear. Certainly, the
Chlorella in a green hydra are immune to being eaten
by filter feeders, have an assured source of mineral nutrients, CO2, and nitrogen, and are moved into light as
the hydra moves. However, the actual rate of increase
of algal cells inside hydra is probably lower than those
of algal cells outside, and it is not obvious that the
number of algal cells contained in the entire green hydra population is a significant fraction of the natural
algal population in the lake or pond.
6. Toxicology
With the development of modern techniques, there
has been an impressive increase in the number and
significance of research programs focused on hydra.
For example, like any small, easily cultured animal,
hydra has been attractive for use in ecotoxicology.
In these tests, the survival or death rate of organisms
is typically related to concentrations of particular
toxins.
Toxicity tests using specially prepared hydra can
provide evidence as to whether or not a particular
toxin is likely to cause cancer. The procedure involves
mass culture of Hydra attenuata, and disassociation of
the hydra into individual cells that are aggregated into
“artificial embryos.” The small, mixed-cell hydra patties organize themselves into complete and perfect animals in less than a week. It has been found that toxins
which do not produce cancers are approximately
equally toxic to intact hydra and to the reconstituting
patties while known carcinogens are much more toxic
to the regenerating cell mass than to the intact hydra
(Johnson et al., 1982).
7. Molecular Biology
The regeneration studies, and the toxicological
studies as well as mathematical analyses of hydra population dynamics and evolution (Slobodkin et al.,
1991), are all possible because of the extreme anatomical and behavioral simplicity of hydra.
Loewenhoek first described hydra in 1704. They
became an object of intense scientific concern with the
work of Trembley (Lenhoff and Lenhoff, 1986). Trembley collected and described green and brown hydra,
feeding them on small invertebrates and dissecting
them with a razor. He established their remarkable
capacity to recover from damage that would kill most
animals. Among the ingenious dissections were
removal and splitting of the head, and producing multiheaded animals. He also sliced hydra into rings. In
145
those cases, a new head developed if the slice had been
nearer the original head, while a foot developed if the
slice had been closer to the original foot. He even
threaded whole hydra onto bristles and watched as
they turned themselves inside out, leaving the bristle
behind. There existed in the 18th century a belief that
animals at birth were endowed with a kind of uniqueness, an animal soul. Transforming a single hydra into
10 others identical with itself by slicing it into pieces effectively eliminated the idea of a unique animal soul
(Burnett, 1973).
It was also generally assumed in those days that
green organisms that benefited from light were plants.
Trembley demonstrated that green hydra were animals
that in certain respects acted as if they were plants.
Studies of hydra continued to contribute to our understanding of basic biological questions through the
next two hundred years. Hydra research has gained in
importance in the past two decades since it has been
found that the molecular and genetic apparatus of hydra is also amenable to analysis. The current volume of
work is very large; hence, only a brief indication of
what is being done will be included here.
The biochemical and genetic properties of Cnidaria
are of significance in studies of the evolution of metazoa. Some early workers felt that hydra, arguably the
simplest metazoa, must be very close to the ancestor of
all metazoa. It is now clear that neither hydra itself nor
other cnidarians, such as Scyphozoa, Cubozoa, and
Hydrozoa (to which hydra belongs), are on the main
branch that led from Cnidaria to other metazoa. This
conclusion is based on the fact that they are the only
metazoa in which mitochondrial DNA is arranged in a
linear, rather than circular pattern. However, another
major group of Cnidaria, the anthozoans, have circular
molecules of mitochondrial DNA like all other metazoa
and would, therefore, seem closer to the main line of
metazoan evolution than hydra (Bridge et al., 1992;
Warrior and Gall, 1985).
Despite this finding, the extreme simplicity of hydra anatomy suggests that it may provide a useful molecular model for higher metazoa. Signal transduction
pathways mediated by cAMP or inositol triphosphate
are present in hydra as they are in virtually all higher
organisms.
The control of basic body form in hydra involves
an interaction of homeotic genes, some of which are
largely identical with body form genes in mice and fruit
flies (Bernfield, 1993; Aerne, 1995; Finnerty and
Martindale, 1999; Galliot et al., 1999). The implications of this are that the basic developmental processes
of the first metazoa (or perhaps advanced premetazoan ancestors of the first metazoa) evolved a
developmental control mechanism which has remained
146
Lawrence B. Slobodkin and Patricia E. Bossert
FIGURE 9 (A) Craspedacusta sowberii, also named Limnocodium victoria (see text); (B) larval stage of A,
also named Microhydra ryderi (see text); and (C) stages in development of Craspedacusta [redrawn from
Payne, 1924.]
essentially intact throughout all subsequent metazoan
evolution!
B. Craspedacusta
The first scientific accounts of freshwater medusae
were based on specimens found in the giant water lily
tank of Regents Park in London in 1880 (Fig. 9A).
Two authors described these separately. Lankester
named them Craspedacusta sowberii after the discoverer, Sowber, and Allman named the same organisms
Limnocodium victoria after the lily. In the same year
and in the same tanks, a tiny colonial hydroid was
found. This animal had no tentacles; each polyp terminated in a bulbous “capitulum” studded with nematocysts. In the center of the capitulum was the mouth.
These little polyp colonies (Fig. 9B) were initially, and
correctly, assumed to be the larval stage of the
medusae. However, very similar polyps were discovered
in a water tank in Philadelphia and were described and
named as a separate species. It was not until 1928 that
it became once again obvious that the polyp named
Microhydra ryderi was the larva of Craspedacusta
(Boulenger and Flower, 1928).
Since their initial description, Craspedacusta
medusae have been found in many locations around
the world, apparently transported with ornamental
aquatic plants and with the water hyacinth, a recently
spread pest species. They have also been found in locations free of imported plants; for example, Lake Gatun
in the Panama Canal zone (Slobodkin, personal observation). Since first documented in the United States in
1908, these unpredictable and sporadic organisms have
been reported in 35 states. The jellyfish have been
5. Cnidaria
found in lakes, farm ponds, quiet coves of rivers, and
old water-filled quarries. When they occur they are
locally abundant but sufficiently rare to call to public
attention (Anonymous, 1999).
1. Life Cycle
The most conspicuous stage of the life cycle is the
small medusa (0.5 – 1.5 cm). Except in the Yang-tse
river system of China, the occurrence of noticeable
populations of the medusae of Craspedacusta is sporadic and often surprising to local naturalists. As the
water warms up in a pond or in the slow current of a
stream backwater, a swarm of medusae appears, often
where it has never been seen before or at least has not
been apparent for many years. These medusae feed on
zooplankton. As they grow, the number of tentacles increases from 8 – 12 up to as many as 100. After several
weeks of growth, gonads develop in pouches of the radial canals. Fertilized eggs produce a small crawling
planula, which then differentiates into a microhydra
(Payne, 1924). The planula consists of two cell layers
forming a double-walled, sausage-shaped sac. The differentiation of the planula into a microhydra is rather
simple. It stands on end and both a mouth and a capitulum develop on the unattached end, thereby forming a
polyp.
The new polyps may continue to bud off new microhydra or medusae. Buds may remain attached after
they have developed a capitulum, producing colonies of
up to 12 polyps attached to a common stolon. Starvation and severely abnormal temperatures cause the
polyps to shrink to a cellular ball, surrounded by a
chitinlike membrane. This can persist through the severe conditions and then redifferentiate as a polyp.
The buds of medusae appear initially as rounded
swellings of the polyp wall (Fig. 9C). Over a period of
several weeks, they enlarge and develop a central
manubrium. An endoderm-lined circular canal with
four radial canals leads to the base of the manubrium.
Eight tentacles emerge from the circular canal. The
medusa bud is initially covered by a layer of tissue that
eventually perforates centrally, remaining as the vellum
of the adult. The mouth opens into the manubrium.
The medusa is by now at least as large as the polyp, to
which it is still attached at the aboral end. Eventually,
it begins locomotory pulsation and separates from the
polyp, completing the life cycle (Payne, 1924).
Medusae usually reproduce sexually; fertilized eggs develop into planulae, which transform into microhydra.
2. Ecology
The microhydra polyps feed in essentially the same
fashion as hydra. Their small size limits their prey, but
this is partially compensated for by their colonial
147
growth pattern, which permits several polyps to make
a simultaneous attack. Free-living medusae feed on various crustacean zooplankters. They can even kill, but
apparently not swallow, the large (0.5 cm), predaceous
cladoceran Leptodora (Dodson and Cooper, 1983).
As a medusa feeds, it grows larger, adding additional tentacles at both the vellum margin and its inner
edge. Several hundred tentacles are found on mature
animals. Medusae appear sporadically in shallow
ponds, natural lakes, and artificial reservoirs throughout the north and south temperate zones. Often entire
medusa populations are unisexual. The polyps are so
inconspicuous that they probably have a much broader
distribution than the literature reports.
Many locations containing polyps are reported not
to have medusae. This bewildering picture has been
clarified by Acker (1976) and Kramp (1950) . Acker
suggested that medusa production by the polyps requires temperatures greater than approximately 20°C
during a period of increasing temperature and adequate, but not enormous, food levels. Other environmental alterations may be significant but have not been
tested. These conditions are reminiscent of those that
produce sexuality in hydra.
Acker and Kramp maintain that the original natural habitat of Craspedacusta sowerbii is the Yangtse-Kiang region of China. In the upper river valley,
two Craspedacusta species, C. sowerbii and C. sinensis,
coexist, whereas only C. sowerbii reaches the downstream areas. From this habitat, C. sowerbii traveled
with water hyacinths and other plants to its present
worldwide distribution.
Shallow pools exist along the lower Yang-tse valley
which are subject to large temperature changes and to
sudden flooding from the main river during high water.
Plankton populations in these ponds fluctuate strongly.
In these ponds, medusae are a recurring phenomenon
throughout the year. The occurrence of medusae in the
spring is so regular that they are given a common name
that translates to “peach blossom fish.”
The generic name Limnocodium is usually applied
to freshwater medusae of the Old World and Craspedacusta to those of the New; but it is not clear if this is
more than a geographic distinction nor has there been
enough investigation to determine how many species of
freshwater medusae actually exist. Due to their sporadic occurrence, the changes in size and tentacle number with developmental stage, the simplicity of larval
anatomy, and the difficulties of preserving specimens,
morphologic studies of freshwater medusae are difficult. As in hydra, many species have been described,
but perhaps some of these are based on nutritional or
developmental history or preservation artifacts. The
strongest evidence for multiple species is that sympatric
148
Lawrence B. Slobodkin and Patricia E. Bossert
populations of two species of medusae are known from
the Yang-tse and that a medusa from Lake Tanganyika
has an extraordinarily different life cycle. In this
latter case, a mature medusa buds new medusae from
its manubrium in addition to reproducing sexually
(Bouillon, 1957).
C. Calposoma
This is a colonial polyp not much bigger than a
paramecium and similar in general appearance to the
microhydra except that it is considerably smaller and
has tentacles rather than a capitulum. As far as is
known, there is only one species, Calposoma dactyloptera (Fuhrman, 1984). Some of its properties have
been described by Rahat and Campbell (1974). These
organisms are so small that their tentacles consist of a
single cell, a tentaculocyte, which contains a row of
miniscule nematocysts. The full natural history of these
animals and the details of their life cycle are not
known. Their general anatomy is reminiscent of a
miniaturized hydra with stiff tentacles, but this does
not necessarily indicate taxonomic or evolutionary
proximity.
D. Cordylophora
The genus Cordylophora is an athecate member of
the primarily brackish water and marine hydrozoan
family Clavidae. It grows as a branching colony up to
5 cm high. The feeding polyps have a conical hypostome
on which filiform tentacles are irregularly arranged.
Colonies also include gonophores, which produce gonads. There is a chitinous periderm (Fig. 10). During
periods of stress, the animals regress to “metanonts”,
masses of resting tissue in the hydrorhizae (Naumov and
Stepan’iants, 1980). The metanonts appear when the
plant stalks on which they grow begin decomposing in
the fall (Roos, 1979).
Cordylophora was first found in the Caspian and
Black seas. It has been suggested that it evolved during
the time that these two bodies of water were connected. It now occurs in rivers of Europe and America,
apparently having spread during the last century attached to ship bottoms. The recent expansion of range
may be attributed to the relatively greater speed of
seagoing vessels during the last hundred years which
would shorten the time of salt water immersion between brackish water ports (Roos, 1979).
Industrial and navigational developments have
extended the region of brackish water in some estuaries, and the increasing pollution of rivers may have
duplicated estuarine conditions. This may be expected
to encourage the geographic spread and the upriver
FIGURE 10
Cordylophora [Redrawn from Roos, 1979].
movement of not only Cordylophora but also other estuarine Cnidaria (Hubschman and Kishter, 1972). The
possibility of foreign Cnidaria appearing in habitats
where they have never been seen is also enhanced by
rapid vessels and the use of water ballast discharged in
ports of arrival. Cordylophora is one example of an expanding range. We expect that there are many others,
which we have not attempted to survey.
E. Polypodium Hydriforme
The remaining freshwater coelenterate that we will
consider is the strange and poorly understood Polypodium hydriforme, originally described by (Lipin,
1911, 1926) and later examined in detail by Raikova
(1973, 1980). This tiny Hydrozoan was first discovered
in Eastern Europe as a parasite inside the eggs of the
European sterlet (Acipenser ruthenus), a small member of the sturgeon family. It is particularly interesting
as it is perhaps the only endoparasitic coelenterate.
5. Cnidaria
149
FIGURE 11 (A) Polypodium stolon with apex of knobs directed toward center of egg; (B) emerging polyp;
and (C) mature polyp [redrawn from Lipin, 1911.]
Obviously there are modifications associated with its
parasitic habit that make classification difficult, but it is
reported to have only one type of nematocyst. These
match the type found in the narcomedusae, an extremely toxic group of hydrozoans.
The development of the larval Polypodium
(Fig. 11) is closely coordinated with that of the sturgeon egg. A binucleate, single cell stage is known from
immature sterlet oocytes. The two nuclei are unequal in
size and chromosome number. The smaller, reportedly
haploid nucleus is surrounded by the large polyploid
nucleus, which develops into a trophic envelope around
the embryo formed by the division of the small nucleus
and its surrounding cytoplasm. By the time the host
oocyte has started to accumulate yolk, the Polypodium
is a two-layered planula, approximately 1-mm long. It
has a flagellated external layer, which eventually will
become the endoderm and an internal layer of ultimate
ectoderm, all surrounded by a capsule that serves as a
digestive organ for consuming yolk (Raikova, 1980).
After a month, the planula has developed into a
stolon with internally directed buds and tentacles. At
this stage, the Polypodium is a colony consisting of a
straight stolon from which project as many as a dozen
knobs, arranged linearly with their apices toward the
center of the egg (Fig. 11A). Each knob develops two
indentations. From each indentation, 12 tentacles, four
of which are short and stubby, project into the stolon
itself (Fig. 11B).
As the fish eggs ripen and are released from the
fish, the tentacles evert through a slit in the stolon.
Simultaneously, the knobs invert, developing a coelenteron lined by what had been the surface exposed to
the egg yolk. The stolon breaks up and the knobs now
appear as somewhat bifurcated polyps with 12 tentacles on each head and a coelenteron full of fish egg yolk
(Fig. 11C).
The free-living polyps subdivide by longitudinal
fission. They crawl on the bottom using tentacles as
walking legs, aided by nematocysts (isotrichous
isorhiza) that hold the substrate. They feed on turbellaria and oligochaetes. Gonads form on the polyps:
both single sex and hermaphroditic individuals are
known. The genital anatomy is considerably more
complex than that of hydra. These gonads and their accessory structures arise from endoderm. The presumed
ovule is diploid and is released into the gastric cavity.
The presumed male gonads become filled with binucleate cells. One nucleus remains haploid, while the other
becomes polyploid. These may fall out of the coelenteron, but there are observations of polyps crawling
onto young sterlets and placing these gonads on the
fish. The transition from fish surface to immature
oocyte has not been observed.
Polypodium parasitizes at least five species of
Acipenser in all of the major rivers of the former Soviet
Union. Eighty percent of sterlet (Acipenser ruthenus)
and 20% of the sturgeon (A. guldenstadti) are infested,
150
Lawrence B. Slobodkin and Patricia E. Bossert
with sporadic infections in other species of the genus. A
careful search of North American sturgeon will probably reveal the presence of this cnidarian.
IV. COLLECTION AND MAINTENANCE
OF FRESHWATER CNIDARIA
A. Collecting Techniques
Hydras are so ubiquitous that their presence
should be expected in any reasonably unpolluted body
of water. Unfortunately, since they are usually sedentary, they are not easily found in plankton tows. Also,
lacking hard body parts, they are often badly damaged
by preservatives. A careful examination of suitable substrates is required. If rocks, leaves, submerged vegetation (e.g., Myriophyllum, Elodea) are collected and
placed overnight in a glass or enamel pan, and the pan
is carefully examined under a low-power dissection
microscope, hydras are usually found. Abundance will
depend on the seasonal distribution of zooplankton
and may vary among lakes.
Since hydras float when hungry and attach again
after they have been fed, the best place to took for
them is often the downstream end of a lake or the
pools in the stream immediately below the lake. If there
is a dam, this should be examined with particular care.
In a typical body of water, there may be at least three
species of Hydra: a green one, a large brown one, and
an intermediate-sized brown one.
Collecting of the medusae of Craspedacusta must
be done with buckets rather than nets. The greater the
sample volume the better (to prevent anoxia and excess
temperature change). A compromise must be found
between excessive agitation of the water, which will
damage the animals, and insufficient stirring, which
will permit them to settle or to become anoxic. The
microhydra larvae can be found on fragmentary
organic debris, much like hydra, but we have not collected them.
B. Maintenance Procedures
Craspedacusta and Cordylophora can be kept in
natural water and fed on field-collected zooplankton.
Even the free-living stage of Polypodium can be kept in
the laboratory and fed on oligochaetes and turbellaria
(Raikova, 1973). Dodson and Cooper (1983) have fed
a range of foods to Craspedacusta, including large rotifers and copepods.
Generally, it is extremely difficult to maintain jellyfish in aquaria for any length of time. Their tissues are
so fragile that they are battered to pieces by most
aquarium aerating or stirring systems. The more or
less sedentary polyps are therefore easier to study.
However, the microhydra larvae of Craspedacusta
have not been extensively studied, perhaps due to their
small size.
Hydras are laboratory animals par excellence. General directions for most of the things one might want to
do with hydra are discussed in detail in the papers collected by Lenhoff (1983). They can be maintained in
artificial pond water or bottled spring water and fed on
either live natural foods, such as cladocerans (e.g.,
Daphnia), copepods, or larvae of midges and mosquitoes. They will not eat protozoa.
Although the brine shrimp, Artemia, do not normally occur in freshwater, many pond animals eat them
avidly in the laboratory. Many investigators use
Artemia nauplii as food for hydra, Craspedacusta, planarians, and other small freshwater invertebrates. The
Artemia eggs are collected at the edges of salt ponds
and lakes. They are sold in vacuum-packed cans for use
by aquarists and may be stored for long periods at cool
temperatures.
The procedure for feeding hydra with Artemia consists of following the package directions for hatching
the brine shrimp (a process requiring about 1 day),
draining and rinsing the hatched nauplii free of the salt
water, and adding them to the hydra. The nauplii must
be alive and vigorous or the hydra will not eat them.
Dead Artemia can be fed to flatworms! Artemia are
best drained in a net that can be made by inserting a
taut sheet of bolting silk in an embroidery hoop. The
Artemia and their salt solution are poured into the net,
which should be wet on both sides to speed drainage.
The shimmering mass of Artemia and eggs are then
rinsed into a new container of hydra culture water. If
this is permitted to stand for 15 min, the active nauplii
will swim toward a light, leaving the unhatched eggs
behind. These active swimmers can be taken with a
medicine dropper and fed to the hydra. This keeps salt
and hydra separate and also permits elimination of
most of the unhatched Artemia eggs.
These small prey may be eaten in great numbers.
Large brown hydra can eat as many as 100 Artemia at
a single meal, and even the smallest green hydra can
consume 1 or 2. No more than 10 min after the first
prey is swallowed, the swallowing process stops even if
some prey remain on the tentacles. We have seen that
organisms may brush against the tentacles with impunity for some time after a hydra has been satiated.
This suggests either that nematocysts are no longer discharging or that the nematocyst supply has been temporarily exhausted.
The primary danger in this feeding process is that
water contaminated with either dead Artemia or with
5. Cnidaria
the regurgitation products of previous feedings is
highly detrimental. The live food must therefore be
added to the hydra; and after approximately 1 h, the
hydra must be placed in clean water, either by moving
them or by discarding the tainted water. Since hydra
usually stick to the substrate, the old medium can be
simply poured off and a new medium added. The glass
surface will eventually become coated with bacteria.
This can be prevented somewhat by scraping the glass
with a rubber spatula before discarding the old
medium, but this is not usually effective for more than
about 1 week, at which time it is best to provide new,
clean containers. Hydras are remarkably sensitive to
heavy metals and to detergents. Even a very short
length of copper tubing in a water supply will kill
hydra. Therefore, in cleaning the dishes, physical dirt is
often less dangerous than the detergents. Dishes must
be very thoroughly rinsed in metal-free water. To obtain nontoxic water, one may either use water from the
collecting site, pretest the water with hydra to make
sure that the water is innocuous, or use glass distilled
or deionized water. When rinsing dishes containing hydra, appropriate salts must be added to the distilled
water.
Various recipes for culture media are available. We
use two stock solutions adapted by K. Dunn (personal
communication) from solutions developed by Loomis,
Lenhoff, and others. Solution A contains 81 g NaHCO3
in 1 L of distilled water. Solution B contains 7.46 g KCl,
20.33 g MgCl2 6H2O, and 147.02 g CaCl2 2H2O in
1 L of distilled water. These stock solutions are added at
the rate of 1 mL/L to distilled water to make artificial
pond water. Hydras die at temperatures above approximately 30°C, but most seem healthy at temperatures as
low as 5°C. The growth and reproductive rates are proportional to temperature, as are the food demands.
Stocks are therefore best maintained at low temperatures except when rapid growth is desired.
V. CLASSIFICATION OF FRESHWATER CNIDARIA
The five groups of freshwater Cnidaria are clearly
distinct from each other. Except for the hydras, they
are relatively rare and sporadic in their distribution. In
North America there is no evidence, at present, for
more than one species of Craspedacusta, Polypodium,
or Calposoma. There may be several species of Cordylophora, but we are considering Cordylophora as one
example of a brackish water cnidarian rather than a
completely freshwater coelenterate. Our taxonomic
key, therefore, consists of five very coarse divisions, one
of which, that for the hydras, is then subdivided in
somewhat greater detail.
151
A. Species Groups of Hydras
There are two closely related genera of hydras: the
brown genus known as Hydra and the green hydras assigned to the genus Chlorohydra. By convention, the
term hydra refers to members of both genera, unless
otherwise specified. Hydras tend to look superficially
similar, except for the distinction between the asymbiotic and green species. On more careful examination,
differences become apparent, not only among apparently unrelated species but even within a clone. Asymbiotic animals from the same clone change color depending on the color of their food. All hydra will also
change body size as a function of the amount of food
and of temperature (Hecker and Slobodkin, 1976). Despite the within-clone plasticity, unrelated hydra maintained under identical conditions, side by side in the
laboratory, retain consistent differences. Batha (1974)
noticed that field-collected hydra differ in appearance
from their own clonal descendants maintained for long
periods in the laboratory. Even animals collected from
the same pond at different seasons and kept in the laboratory sometimes show consistent differences in laboratory cultures, despite apparent similarities of the
field-collected specimens. The effect of all this is to
make taxonomy extremely difficult. Some investigators
have tended to describe new species on the basis of
rather unstable, variant appearances, while others have
despaired of making any precise identifications.
Campbell (1987) has provided a compromise position, in which he has classified the hydra into clearly
distinct “species groups,” each composed of an uncertain number of more closely related species whose precise identity may have to await new techniques of
examination. Color, presence or absence of a body
stalk, nematocyst shape, and the order of appearance
of tentacles on the new buds are the characters used.
The structure of the theca around the fertilized eggs,
the appearance of gonads, proportions of different
kinds of nematocysts, microscopic details of symbiotic
algae, and certain physiological characteristics are all
likely to be important in subdividing the various
species groups. Unfortunately, these subdivisions are
difficult for all but the most serious students and professionals. Since the species group is adequate for most
purposes, it seems advisable to quote extensively from
Campbell’s descriptions. The four species groups are
given as follows.
1. The easily recognized group Hydra viridissima
(also known as Chlorohydra). In addition to the
brilliant green color, Campbell lists the following characteristics: “small to moderate body
size and tentacles, stalk not distinct from the
rest of the column; hermaphroditic;
152
Lawrence B. Slobodkin and Patricia E. Bossert
FIGURE 13 (A) Stenotele and (B) tentacle formation in Hydra oligactis group. [Redrawn from Campbell, 1987.]
FIGURE 12
(A) Holotrichous isorhiza and (B) nonspiny embryotheca. [Redrawn from Campbell, 1987.]
embryotheca spherical, made of polygonal
plates; nematocysts very small; tentacles arising
simultaneously on buds.”
2. The stalked Hydra or Oligactis group: “large
size; pronounced translucent stalk in large
individuals; long tentacles; dioecious; spherical
embryo with simple theca; slender stenotele
(capsules) and large, blunt, cylindrical holotrichous isorhiza (capsules); distinct golden color in
culture due to yellow crystals in the ectoderm;
two lateral tentacles arising before the others in
the bud” (see Fig. 12).
3. The vulgaris group or common hydra “11 moderate size tentacles of moderate length; dioecious or monoecious, sometimes switching
between the two conditions; spherical
embryotheca ornamented with spines; broad
stenotele (capsules); slender holotrichous
isorhiza; often slipper-shaped, with the anterior
end broadly pointed; tentacles arising on the
buds simultaneously or nearly so” (see Fig. 13).
4. The “gracile Hydra” or braueri group: “small
to medium size; tentacles moderately short; hermaphroditic; embryotheca and egg flattened,
with the embryotheca adherent to the substrate;
embryotheca smooth or papillate; tentacles arising simultaneously on the buds; holotrichous
isorhiza (capsule) broader than half its length;
and stenotele (capsule) plump; body often
palecolored during laboratory cultivation.”
No single character, not even the nematocysts, can
by itself distinguish one group from another. Every
group probably contains a large number of species,
each of which has been described under a large number
of names. Perhaps biochemical procedures may eventually sort them out, but for the moment and for most
purposes, strains collected from the field should be
classified to a species group and then supplied with
enough ancillary description so that other workers can
at least know if they have the same or a similar organism. Campbell (1987) provided a key to the four
groups of Hydra, which we have incorporated in a key
to the other freshwater Cnidaria.
B. Taxonomic Key to Genera of Freshwater Cnidaria
The polyp and medusa stages of a single species of
coelenterate can differ greatly, both morphologically
and ecologically. The key has been arranged to give the
same results whether one starts with polyps or
medusae. Also, the key refers to “species groups” of
Hydra as names in parentheses.
1a.
Parasitic in fish eggs ..........................................................................................................................................................Polypodium
1b.
Nonparasitic ......................................................................................................................................................................................2
2a(1b).
Medusae ........................................................................................................................................................................Craspedacusta
2b.
Polyps ................................................................................................................................................................................................3
3a(2b).
Polyp without basal attachment, with oral surface downward ..........................................................................................Polypodium
3b.
Polyps with basal disk ........................................................................................................................................................................4
4a(3b).
Solitary polyps with single circle of multicellular tentacles; no medusa, gonads are body stalk...........................................................7
5. Cnidaria
153
4b.
Colonial polyps ..................................................................................................................................................................................5
5a(4b).
Filiform tentacles, irregularly arranged on conical hypostome; branching colony with gonophores ...............................Cordylophora
5b.
Colonial; atentacular or tentacles unicellular......................................................................................................................................6
6a(5b).
Atentacular, oral capitulum; polyps from common stolon; may bud medusa from polyp;
“microhydra” ................................................................................................................................................................Craspedacusta
6b.
Minute with unicellular tentacles ........................................................................................................................................Calposoma
7a(4a).
Bright green...................................................................................................................................................................Hydra (viridis)
7b.
Not green ...........................................................................................................................................................................................8
8a(7b).
Holotrichous isorhiza nematocysts at least half as broad as long; embryotheca without
spines...........................................................................................................................................................................Hydra (braueri)
8b.
Holotrichous isorhiza nematocysts slender or if plump, then embryotheca spined..............................................................................9
9a(8b).
Buds acquire two lateral tentacles before others appear; otherwise stenotele nematocysts at least 1.5 times as long as
broad .........................................................................................................................................................................Hydra (oligactis)
9b.
Buds acquire tentacles in some other order; otherwise stenotele nematocysts less than 1.5 times as long as broad
...................................................................................................................................................................................Hydra (vulgaris)
LITERATURE CITED AND FURTHER READINGS
Acker, T. S. 1976. Craspedacusta sowerbii: an analysis of an introduced species, in: Mackie, G. Ed., Coelenterate biology and behavior, Plenum, New York, pp. 219 – 226.
Aerne, B. 1995. Cnidarian hox genes. Developmental Biology
169:547 – 56.
Angradi, T. R. 1998. Observations of freshwater jellyfish, Craspedacusta sowerbii Lankester (Trachylina: Petasidae), in a West
Virginia reservoir. Brimleyana 25:34 – 42.
Anonymous. 1999. Fresh water jellyfish in Rhode Island, URI Watershed Watch Home Page.
Barnes, R. D. 1966. Invertebrate Zoology, Saunders, Philadelphia, PA.
Batha, J. V. 1974. The distribution and ecology of the genus Hydra
in the Milwaukee area of Lake Michigan. Ph. D. thesis, Zoology Department, University of Wisconsin, Milwaukee.
Bernfield, M. 1993. Molecular basis of morphogenesis. [Symposium
with the Society for Developmental Biology] Wiley-Liss, New
York.
Bossert, P. 1986. Regulation of intracellular algae by various strains
of the symbiotic Hydra viridissima. J. Cell Sci. 85:187 – 195.
Bossert, P. 1988. The effect of Hydra strain size on growth of endosymbiotic alga. Ph.D. thesis, Department of Biology, State
University of New York, Stony Brook, NY.
Bossert, P., Dunn, K. 1996. Regulation of intracellular algae by various strains of the symbiotic Hydra viridissima. Journal of Cell
Science 85:187 – 195.
Bouillon, J. 1957. Etude monographique du genre Limnocnida (Limnomedusae). Annales de la Societe Royale Zoologique de Belgique 87:254 – 500.
Boulenger, C. L., Flower, W. U. 1928. The Regent’s Park medusa,
Craspedacusta sowerbii and its identity with C. (Microhydra)
ryderi. Proceedings of the Zoological Society of London
66:1005 – 1015.
Bridge, D., Cunningham, C., Schierwater, B., DeSalle, R., Buss, L.
1992. Class level relationships in the phylum Cnidaria: Evidence from mitochondrial genome structure. Proceedings
National Academy of Science USA 89:8750 – 8753.
Burnett, A. L. 1973. Biology of Hydra, Academic Press, New York.
Campbell, R. D. 1987. A new species of Hydra (Cnidaria: Hydrozoa)
from North America with comments on species clusters within
the genus. Zoological Journal of the Linnean Society
91:243 – 263.
Campbell, R. D. 1999. The Hydra of Madagascar (Cnidaria: Hydrozoa). Annales de Limnologie-International Journal of Limnology 35(2):95 – 104.
Ciernichiari, E., Muscatine, L., Smith, D. C. 1969. Maltose excretion
by the symbiotic algae of Hydra viridis. Proceedings of the
Royal Society of London Series B 173:557 – 576.
Colasanti, M. 1995. NO in hydra feeding response. Nature
374(6522):505.
Devries, D. R. 1992. The freshwater jellyfish Craspedacusta sowerbyi: a summary of its life history, ecology, and distribution.
Journal of Freshwater Ecology 7(1):7 – 16.
Dodson, S. I., Cooper, S. D. 1983. Trophic relationships of the freshwater jellyfish Craspedacusta sowerbii Lankester 1880. Limnology and Oceanography 28:345 – 351.
Dunn, K. W. 1986. Adaptations to endosymbiosis in the green
Hydra, Hydra viridissima. Ph. D. thesis, Department of Biology,
State University of New York, Stony Brook.
Dunn, K. W. 1987. Growth of endosymbiotic algae in the green
hydra, Hydra viridissima. Journal of Cell Science 88:571 – 578.
Dunn, K. W. 1988. The effect of host feeding on the contribution of
endosymbiotic algae to the growth of green hydra. Biology
Bulletin 175:193 – 201.
Elliott, J. K., Elliott, J. M., Leggett, W. C. 1997. Predation by hydra
on larval fish: field and laboratory experiments with bluegill
(Lepomis macrochirus). Limnology and Oceanography 42(6):
1416 – 1423.
Finnerty, J. R., Martindale, M. Q. 1999. Early evolution of Hox and
ParaHox genes: evidence from the Cnidaria. Developmental
Biology 210(1):41.
Fuhrman 1984. Craspedacusta sowerbii Lank. et un noveau coelentere d’eau douce, Calpasoma dacryloprera. n.g., n.sp. Revue
Suisse de Zoologie 46:363 – 368.
Galliot, B., de Vargas, C., Miller, D. 1999. Evolution of homeobox
154
Lawrence B. Slobodkin and Patricia E. Bossert
genes: Q(50) Paired-like genes founded the Paired class. Development Genes and Evolution 209(3):186 – 197.
Griffing, T. C. 1965. Dynamics and energetics of populations of
brown hydra. Ph.D. thesis, Zoology Department, University of
Michigan, Ann Arbor, MI.
Grosvenor, W., Rhoads, D. E., Kassimon, G. 1996. Chemoreceptive
control of feeding processes in hydra. Chemical Senses
21(3):313 – 321.
Hecker, B., Slobodkin, L. B. 1976. Responses of Hydra oligactis to
temperature and feeding rate, in: Mackie, G. O. Ed., Coelenterate ecology and behavior, Plenum, New York, pp. 175 – 186.
Hershey, A., Dodson, S. 1987. Predator avoidance by Cricotopus: cyclomorphosis and the importance of being big and hairy. Ecology 68:913 – 920.
Hessinger, D. Lenhoff, H. M. Eds. 1988. The biology of nematocysts,
Academic Press, London.
Hubschman, J. H., Kishler, W. J. 1972. Craspedacusta sowerbii
Lankester 1880 and Cordylophora lacustris Allman 1871 in
Western Lake Eric (Coelenterata). Ohio Journal of Science
72:318 – 332.
Hyman, L. H. 1940. The invertebrates: Protozoa through
Ctenophora, McGraw-Hill, New York.
Johnson, E. M., Gorman, R. M., Gabel, B., George, M. E. 1982. The
Hydra attenuata system for detection of teratogenic hazards.
Teratogenesis, Carcinogenesis, and Mutagenesis 2:263 – 276.
Kanaev, I. I. 1969. Hydra: essays on the biology of freshwater polyps.
Lenhoff, H. M. Ed. [Translated by Burrows, E. T. and Lenhoff,
H. M.] Published by H. T. Burrows and H. M. Lenhoff,
University of Miami, Coral Gables, FL. [Originally published by
the Soviet Academy of Sciences, Moscow, 1952.] 453 pp.
Kouchinsky, A., Bengtson, S., Gershwin, L. A. 1999. Cnidarian-like
embryos associated with the first shelly fossils in Siberia. Geology 27(7):609 – 612.
Kramp, P. L. 1950. Freshwater medusae in China. Proceedings of the
Zoological Society of London 120: 165 – 184.
Lenhoff, H. M. 1974. On the mechanism of action and evolution of
receptors associated with feeding and digestion, in: Muscatine,
L. Lenhoff, H. M. Eds., Coelenterate biology: reviews and new
perspectives, Academic Press, New York, pp. 211 – 214.
Lenhoff, H. M. 1983. Hydra: research methods, Plenum Press, New
York, 463 pp.
Lenhoff, H. M., Loomis, W. F. 1961. The biology of Hydra and of
some other coelenterates, University of Miami Press, Coral
Gables, FL.
Lenhoff, S. G., Lenhoff, H. M. 1986. Hydra and the birth of experimental biology, 1744: Abraham Trembley’s Memoires concerning the polyps, Boxwood Press, Pacific Grove, CA.
Link, J., Keen, R. 1995. Prey of deep-water hydra in Lake Superior.
Journal of Great Lakes Research 21(3):319 – 323.
Lipin, A. 1911. Morphologie und biologie von Polypodium. Zoologische Jahrbuecher, Abteilung fuer Anatomie und Ontogenie der
Tiere 31.
Lipin, A. 1926. Polypodium. Zoologische Jahrbuecher, Abteilung
fuer Anatomic und Ontogenie der Tiere.
Lomnicki, A., Slobodkin, L. B. 1966. Floating in Hydra littoralis.
Ecology 47:881 – 889.
Mackie, G. O. 1976. Coelenterate ecology and behavior, Plenum,
New York.
Martinez, D. 1997. On senescence in asexual metazoans. Ph.D.
thesis, Department of Ecology and Evolution, State University
of New York at Stony Brook, Stony Brook, NY. 181 pp.
Muscatine, L. 1965. Symbiosis of Hydra and algae. III Extracellular
products of the algae. Comparative Biochemistry and Physiology 16:177 – 192.
Naumov, D. V., Stepan’iants, S. D. 1980. Teoreticheskoe i prakticheskoe znachenie kishechnopolostnykh: sbornik. Leningrad,
Zool. in-t.
Pardy, R. 1983. Preparing aposymbiotic Hydra and introducing symbiotic algae into aposymbiotic Hydra, in: Lenhoff, H. M. Ed.,
Hydra: Research methods, Plenum, New York, pp. 393 – 401.
Payne, F. 1924. A study of the freshwater medusae, Craspedacusta
ryderi. Journal of Morphology 38:397 – 430.
Rahat, M., Campbell, R. 1974. Nematocyst migration in the polyp
and 1 celled tentacles of the minute freshwater coelenterate
Calposoma dactyoptgera. Transactions of the American Microscopical Society 93:379 – 385.
Rahat, M., Sugiyama, T. 1993. The endodermal cells of some brown
Hydra are autonomous in their ability to host or not to host
symbiotic algae: an analysis of Chimera. Endocytobiosis and
Cell Research 9(2 – 3):223 – 231.
Raikova, E. 1973. Life cycle and systematic position of Polypodium
hydriforme Ussov (Coelenterata, a cnidarian parasite of the eggs
of Acipenseridae. Publications of the Seto Marine Biological
Laboratory 20:165 – 174.
Raikova, E. 1980. Morphology, ultrastructure and development
of the parasitic larva and its surrounding trophamnion of
Polypodium
hydriforme
Coelenterata.
Cell
Research
206:487 – 500.
Ritte, U. Z. 1969. Floating and sexuality in laboratory populations
of Hydra littoralis. Ph. D. thesis Zoology Department. University of Michigan, Ann Arbor, MI, 350 pp.
Roos, P. 1979. Two stage life cycle of a Cordylophora population in
the Netherlands. Hydrobiologia 62:231 – 239.
Sabine, A., Hoffmeister, H., Chica, H., Schaller, H. C. 1987. Head
activator and head inhibitor are signals for nerve cell and differentiation in hydra. Developmental Biology 122:72 – 77.
Schummer, M., Scheurlen, I., Schaller, C., Galliot, B. 1992. Hom
Hox Homeobox genes are present in hydra (Chlorohydra
viridissima) and are differentially expressed during regeneration. Embo Journal 11(5):1815 – 1823.
Schwartz, S. S., Hann, B. C., Herbert, D. N. 1983. The feeding ecology of Hydra and possible implications in the structuring of
pond plankton communities. Biological Bulletin (Woods Hole,
Mass) 164:136 – 142.
Slobodkin, L. 1964. Experimental populations of Hydrida. British
Ecological Society Jubilee Symposium. Journal of Animal Ecology 33 (Supplement):131 – 148.
Slobodkin, L. 1980. Growth and regulation of animal populations,
Dover, New York.
Slobodkin, L. Bossert, P., Mattessi, C., Gatto, M. 1991. A review of
some physiological and evolutionary aspects of body size and
bud size of Hydra. Hydrobiologia 216:377 – 382.
Spadinger, R., Maier, G. 1999. Prey selection and diel feeding of the
freshwater jellyfish, Craspedacusta sowerbyi. Freshwater Biology 41(3):567 – 573.
Stiven, A. 1976. The quantitative analysis of the hydra-hydramoeba
host – parasite system, in: Mackie, G.O. Ed., Coelenterate ecology and behavior, Plenum, New York, pp. 389 – 400.
Thiel, H. 1973. Limnocnida indica in Africa. Publications of Seto
Marine Biological Laboratory 20:73 – 79.
Thorp, J. H., Barthalmus, G. T. 1975. Effects of crowding on growth
rate and symbiosis in green hydra. Ecology 56:206 – 212.
Walsh, E. J. 1995. Habitat-specific predation susceptibilities of a
littoral rotifer to two invertebrate predators. Hydrobiologia
313:205 – 211.
Warrior, R., Gall, J. 1985. The mitochondrial DNA of Hydra attenuata consists of two linear molecules. Archives des Sciences
(Geneva) 38:439 – 445.
6
FLATWORMS: TURBELLARIA
AND NEMERTEA
Jurek Kolasa
Department of Biology
McMaster University
Hamilton, Ontario L8S 4K1 Canada
TURBELLARIA
I. Introduction: Status in the Animal
Kingdom
II. Anatomy and Physiology
A. General External and Internal
Anatomical Features
B. Environmental Physiology
III. Ecology
A. Life History
B. Distribution
C. Behavioral Ecology
D. Foraging Relationships, Predators,
and Parasites
E. Population Regulation and Density
F. Functional Role in the Ecosystem
IV. Current and Future Research Problems
A. Dispersal
B. Rhabdoids
C. Endosymbiosis
D. Community Ecology
V. Collecting, Rearing, and Identification
Techniques
VI. Identification of North American Genera
of Freshwater Turbellaria
A. Taxonomic Key to Orders and
Suborders of Turbellaria
B. Taxonomic Key to Catenulida
Turbellaria and Nemertea are common and often
very numerous inhabitants of freshwaters. Even
though more than 200 species of Turbellaria and
three species of Nemertea live in North America,
their ecology and systematics have been less studied
than that of many other common aquatic invertebrates. An obvious reason for this limited attention is
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
C. Taxonomic Key to Macrostomida
D. Taxonomic Key to
Lecithoepitheliata
E. Taxonomic Key to Prolecithophora,
Proseriata, and Tricladida
F. Taxonomic Key to Typhloplanoida
G. Taxonomic Key to Kalyptorhynchia
and Similar Forms
H. Taxonomic Key to Dalyellioida
NEMERTEA
VII. General Characteristics, External
and Internal Anatomical Features
VIII. Ecology
A. Life History and General Ecology
B. Physiological Adaptations
C. Behavioral Ecology
D. Functional Role in the Ecosystem
IX. Current and Future Research
Problems
X. Collection, Culturing, and
Preservation
XI. Taxonomic Key to Species of Freshwater
Nemertea in North America
Literature Cited
Appendix 1. List of North American Species
of Microturbellaria
the difficulty posed by preservation. Most turbellarians become unrecognizable after a routine preservation of field samples in alcohol or formalin. Study of
live specimens is the best method and many interesting discoveries lie ahead in the areas of reproductive
biology, dispersal, endosymbiosis, and community
structure.
155
156
Jurek Kolasa
TURBELLARIA
I. INTRODUCTION: STATUS
IN THE ANIMAL KINGDOM
The turbellarian flatworms are the lowest acoelomate Bilateria with only a single opening to the digestive tract; that is, they lack a definitive anus. Cestoidea
(tapeworms), Trematoda (flukes), and Turbellaria constitute the phylum Platyhelminthes. Because of tradition
and practical considerations, turbellarians are divided
into microturbellarians and macroturbellarians. This division does not reflect phylogenetic relationships, but
rather superficial morphological similarities. Also, depending on the methodology chosen, the taxa discussed
may have a rank different from the one used here (see
Ehlers, 1986). Several orders and suborders of Turbellaria are known from freshwaters (Table I, Fig. 1).
One order, Acoela, has been recently excluded
from Turbellaria on the basis of DNA sequence analysis (Ruiz-Trillo et al., 1999) but we retained it in this
chapter for practical reasons.
Of the approximately 400 species of freshwater
microturbellaria, about 350 have been recorded in
Europe (Lanfranchi and Papi, 1978), and approximately 150 occur in North America (Appendix 6.1),
many of which are the same as those found in Europe.
Many species and genera of microturbellarians remain
TABLE I
to be discovered in North America. Records from other
regions of the world are sparse. Interestingly, Africa,
South America, and Papua New Guinea trail closely
behind North America in the number of species. Differences in species richness are clearly due in great part to
research effort; and for this reason, the number of
species in North America must be highly underestimated. Triclads (often called planarians) are more thoroughly studied. The number of recorded species in
North America is approximately 40, but the actual
number of taxa is likely to be substantially higher.
II. ANATOMY AND PHYSIOLOGY
A. General External and Internal
Anatomical Features
Most freshwater microturbellaria are less than 1
mm in length, although some can reach several millimeters. Triclads are distinctly larger, with most species
exceeding 10 mm and the largest being several centimeters long.
The turbellarian body is elongated, relatively soft,
and usually tapered at the ends. Sometimes lateral flaps
near the cerebral region or a short tail like section are
present. With the exception of triclads, flatworms are
generally not flat — despite their common name. Most
Selected Characteristics of Commonly Recognized Turbellarian Orders and Suborders Occurring in Freshwaters
Taxona
order suborder superfamily
Approx. number of species
Average body size (mm)
Comments and special features
Microturbellarians
Catenulida
60
0.5 – 1
Acoela
Macrostomida
Prolecithophora
Lecithoepitheliata
3
50
5
10
0.5 – 1
1.0 – 3
5.0 – 10
3.0 – 10
Proseriata
Rhabdocoela
Dalyellioida
Temnocephalida
4
2.0 – 5
Thin chains of zooids; common in
various habitats
Most species marine
Common in various habitats
Many marine species
Many marine species; some are
terrestrial or semiaquatic
Many marine species
Typhloplanoida
Typhioplanida
Kalyptorhynchia
Macroturbellarians
Tricladida
a
100
0.8 – 1
1.0 – 14
Common in various habitats
Commensals on crust aceans, snails,
turtles, one parasitie
150
0.5 – 6
15
1.0 – 2
Common in various habitats
including terrestrial and
semiaquatic
Rare; most species marine
100
5 – 20
Taxonomical rank is indicated by indentation.
Greatest diversity associated with
karst habitats
6. Flatworms: Turbellaria and Nemertea
157
FIGURE 1
Schematic representation of higher turbellarian taxa found in freshwaters with an emphasis on
the most useful diagnostic features. (A) Acoela (one unidentified species); (B) Catenulida (e.g., Catenula,
Stenostomum, I and II represent two successive zooids); (C) Macrostomida (Macrostomum, Microstomum);
(D) Lecithoepitheliata (e.g., Prorhynchus, Geocentrophora); (E) Dalyellioida (Gieysztoria, Microdalyellia);
(F) Typhloplanoida (e.g., Mesostoma, Olisthanella, Castrada); (G) Kalyptophynchia (e.g., Gyratrix; intestine
omitted for clarity); and (H) Proseriata and Tricladida (e.g., Bothrioplana, Dendrocoelopsis). Co, Copulatory
organ; cp, ciliated pits; i, intestine, m, mouth; n, protonephridial duct; o, ovary; ph, pharynx; pr, proboscis; s,
stylet, st, statocyst; t, testes; v, vacuolized tissue; and y, yolk glands.
microturbellaria are circular in cross section, with some
differentiation of shape of the dorsal and ventral surfaces. Moreover, species that reproduce asexually may
be composed of several zooids (Figs. 3B, 4A,B, 12A),
giving a chainlike appearance to an animal. Turbellarians may be colorless, white, red, bluish, green, black,
brown, or yellowish depending on epidermal and
parenchymal pigments, gut content, and symbiotic
algae. Some species, such as Dugesia tigrina and
Hydrolimax grisea, develop characteristic patchy patterns of pigments.
Anatomically, the most prominent turbellarian features are a ciliated epidermis, rhabdoids, an intestine
without anus, ventral mouth, and a complex reproductive system (Figs. 1, 9, 12D). The ciliated epidermis
covers the entire body surface. Rhabdoids are rodlike,
light-refracting structures produced in the epidermis
(dermal rhabdoids) or in the parenchyma (adenal rhabdoids). It is often difficult to determine the type of
rhabdoids in live specimens. Adenal rhabdoids are
most abundant at or near the front extremity. They often aggregate in juxtaposed groups that are known in
the taxonomic literature as rod tracts. Wrona (1986)
reviewed hypotheses on the role of rhabdoids in triclads and has provided experimental evidence that
rhabdoids contribute to mucus production. Rhabdoids
are also poisonous and used in prey immobilization or
in deterring predators. Other components of the body
walls include a basal membrane and muscle fibers. The
fibers are longitudinal, circular, or in larger forms, diagonal. Some fibers go through the parenchyma. The
parenchyma is a loose tissue filling the body cavity and
containing various cell types, including secretory,
excretory, reproductive, and formative cells and their
insoluble excretions, that fulfill important physiological
functions (Hyman, 1951). In large turbellarians, the
parenchyma is usually more compact than in smaller
forms, where it can be quite loose and vacuolated.
Turbellarians possess numerous glands, of which the
frontal, pharyngeal, shell-producing, and rhabdoidproducing glands are most common.
A vast array of sensory organs enables turbellarians
to react to environmental stimuli. These organs include
sensory hairs, eyes, statocysts, and chemoreceptors of
various kinds. Some species possess aggregations of sensory cells, or ciliated pits, in lateral depressions situated
in front of the brain ganglion. Exact functions of the
ciliated pits and structures associated with them are
unknown, but at least some of these structures (i.e., the
refractive bodies of Stenostomum) have been implicated
in phototaxis (Marcus, 1951). Some evidence exists that
these refractive bodies function as lenses (Tyler and
Burt, 1988). Thigmotaxis is common and plays a role in
a range of behaviors such as choosing substrate, hunting prey, and avoiding predators. Among North American freshwater Turbellaria, a statocyst is present only
in Catenula, Rhynchoscolex, Otomesostoma, and an
unidentified representative of Acoela (Strayer, 1985;
Kolasa et al., 1987). Turbellarians have two types of
simple eyes: inverted and direct cup eyes (Bedini et al.,
1973). The number of eyes is variable, but one pair of
eyes is most frequently present.
Locomotion in the Turbellaria is based on the
movement of cilia. Some triclads, however, are capable
158
Jurek Kolasa
of fast muscular contractions allowing them to crawl in
bursts of leechlike behavior. Smaller forms can both
swim and glide on submersed objects, while heavier
forms are restricted to gliding only. Adhesive organs in
the form of pseudosuckers or papillae are often present.
B. Environmental Physiology
Gaseous exchange in turbellarians occurs through
their body walls. Such exchange depends on the surface / volume ratio and is not very efficient. Consequently, flatworms may have a limited ability to adapt
to low-oxygen conditions. This conclusion appears to
be true for most flatworms and this limitation is often
exploited in extraction of specimens from samples (e.g.,
Young, 1970; Schwank, 1981). However, the ability to
cope with reduced oxygen levels does not seem to differ
substantially from that of other aquatic invertebrates.
Oxygen adaptability differs more among species
adapted to diverse habitats than among major taxa
(cf. Heitkamp, 1979b). For example, species of
Macrostomum normally found in stream headwaters
are more sensitive to low oxygen levels, and perhaps
high temperature, than are representatives of the
same genus from physically more fluctuating habitats
(J. Kolasa, personal observation). Some species, particularly in the genus Phaenocora, have developed adaptations to cope with anaerobic conditions (Young and
Eaton, 1975). They showed that the presence of algae
in Phaenocora typhlops enhances its ability to survive
low-oxygen conditions. This same species, as well as a
related P. unipunctata (Öersted) produces hemoglobin,
allowing them to store oxygen during burrowing in the
anoxic mud. Many marine species, however, live in
low-oxygen sediments, but have neither hemoglobin
nor symbiotic algae.
Heitkamp (1979a, b) studied respiration rates of
Opistomum pallidum, two species of Mesostoma, and
a species hosting symbiotic zoochlorellae — Dalyellia
viridis. He showed the respiration to be related to the
body size and shape (surface in a very flat Mesostoma
ehrenbergi, or weight in cylindrical Mesostoma lingua
and other species). Interestingly, Dalyellia viridis that
are symbiotic with zoochlorellae had the highest respiration rate of all studied species; possibly their symbiotic algae contribute significantly to the observed rate.
Heitkamp (1979b) found the respiration rate of microturbellarians to vary depending on the temperature and
diurnal rhythm.
A considerable effort has been devoted to studying
nutrition and digestion in Turbellaria, particularly by
Jennings (1977). In this context, the lipid storage
capacity of flatworms and its importance to population
dynamics and competition is discussed for Dugesia
polychroa by Boddington and Mettrick (1977). Specifically, they show that populations are regulated by intraspecific competition for food, which affects individual
size, cocoon production, and cocoon viability.
Triclads clearly avoid light. Microturbellarians appear to be either indifferent or positively phototactic
(e.g., Typhloplana viridata which contains symbiotic
algae), although if a sample is suddenly illuminated,
many will attempt to hide in sediments.
Desiccation of eggs can be tolerated by some
species and it may be an obligatory factor in releasing
them from diapause and stimulating embryonic development. Similarly, some triclads such as Phagocata
may undergo encystment as entire or fragmented animals. Temperature may also have an important effect
on reproduction. Dumont and Schorreels (1990)
showed that clutch size, total offspring, and daily offspring all depended on temperature rather than food in
Mesostoma lingua.
Although the majority of Turbellaria appear to
have some habitat preferences, within a limited range
of physical conditions many species have adaptations
conferring considerable flexibility. Some members of
the Catenulida are found in habitats as different as
fast-flowing streams and temporary pools (e.g., Stenostomum leucops, Catenula lemnae). Mesostoma species
have been reported from both warm rice fields in the
Sacramento Valley in California as well as from snowmelt pools at elevations above 2400 m (Collins and
Washino, 1979).
Even though microturbellarians are delicate and
fragile organisms, they appear to have a significant potential for dispersal. Results of an experiment conducted by Jan Ciborowski (University of Windsor) were
surprising. Ciborowski studied colonization of artifical
ponds (flexible swimming pools) situated on the roof of
a university building for several weeks. About 50% of
these ponds were successfully colonized by six different
species of microturbellaria (unpublished data). With
only one exception, these species are not known to produce resistant disseminules such as eggs or cocoons.
III. ECOLOGY
A. Life History
In most species, miniature replicas of the adult
hatch directly from eggs; these juveniles differ from
adults chiefly by the absence of reproductive systems.
Some freshwater turbellarians, however, may be ovoviviparous (Mesostoma) or may have a larval stage
distinctly different from the adult (Rhynchoscolex).
Numerous modifications of the life cycle have evolved
among turbellarians. Many seasonally occurring species
6. Flatworms: Turbellaria and Nemertea
are univoltine, particularly those associated with
temporary habitats or at the extremes of their geographical ranges. Other species are multivoltine, with
the number of generations depending on habitat availability (see Heitkamp, 1982, for an excellent account).
A similar diversity of life cycles is observed in triclads,
but the ecological reasons for this diversity remain
hidden. Reproductive patterns may sometimes coincide
with major taxonomic subdivisions. Calow and Read
(1986) studied European representatives of the three
families that are also represented in North America:
Dugesiidae, Planariidae, and Dendrocoelidae. These investigators have found that species breeding over several seasons (Dugesiidae, Planariidae) invest less in reproduction than members of Dendroceolidae, which
reproduce once and die.
As a rule, turbellarians are hermaphroditic. Various modifications of both egg formation and development are found and may include self-fertilization, as
in Mesostoma ehrenbergi (Fiore and Ioalé, 1973;
Göltenboth and Heitkamp, 1977), protandry or progyny in many other species, or loss of either the male or
female gonad as in Catenulida (Rieger, 1986). All of
these modifications have implications for habitat
adaptability, clonal diversity, rate of population
growth, and ability to colonize.
Asexual reproduction by means of paratomy, that
is transverse division of the body, is common in several
genera of microturbellaria, particularly in Catenulida
and Macrostomida. In macroturbellarians, architomy
(transversal fission of the caudal region) occurs frequently in Dugesiidae and in some Planariidae (Calow
and Read, 1986; Rieger, 1986). In some species of
Stenostomum, and in some populations of the common
triclad Dugesia tigrina, sexual organs have never been
observed.
B. Distribution
Freshwater turbellarians are largely free-living animals, although a few European freshwater species such
as Varsoviella kozminski (Gieysztor and Wiszniewski)
and Phaenocora beauchampi (Sekera) are ectoparasitic
on crustaceans. Several other freshwater forms, including triclads, occurring in Europe and Australia are
commensal on crustaceans and turtles (e.g., Jennings,
1985). The great majority of freshwater turbellarians
are free-living and live in various aquatic systems such
as ponds, lakes, streams, hyporheic water, ditches, and
temporary puddles. However, some may be found in
aquatic habitats normally excluded from the domain of
freshwater ecology, such as water films among fallen
leaves in a mesic forest or in capillary soil water of a
grassy meadow (Sayre and Powers, 1966).
159
In North America, microturbellarians are known
as far north as Igloolik, a small island at 68°N latitude
off Melville Peninsula. Two species, Mesostoma and an
unidentified mesostomid, were collected by P.D.N.
Hebert and associates (unpublished). In Alaska and in
the area of Hudson Bay in northern Canada, as many
as six different species are known (Holmquist, 1967;
Schwartz and Hebert, 1986). Further south, in temperate climatic zones, the species diversity increases and
can easily reach 20 to perhaps 60 species in a single
lake or pond. For example, there are 23 species of
flatworms in the oligotrophic Mirror Lakes, New
Hampshire (Strayer, 1985) and as many as 57 species
occur in the polytrophic Lake Zbechy in Poland
(Kolasa, 1979). Streams also have rich flatworm fauna.
In a short section of Wappinger Creek, an eastern tributary of the Hudson River in New York, 15 species
were found (Kolasa et al., 1987). A survey of several
streams, including springs and underground water in
the same area revealed 32 species of microturbellarians,
some of which are still undescribed. Because of a
dearth of systematic studies, few microturbellarians are
known from the West Coast. Case and Washino
(1979), Collins and Washino (1979), and other related
papers report Mesotoma lingua from California, together with some other common species in rice fields.
Many species, particularly among the Catenulida
and the Macrostomida, are widely distributed worldwide, while species of the “higher” Turbellaria, such as
the Typhloplanida and Tricladida, are geographically
more restricted. About 3% of all species known from
Europe are designated as cosmopolitan in distribution
(Lanfranchi and Papi, 1978), but studies in the tropics
and Southern Hemisphere (Mead and Kolasa, 1984)
suggest that this figure is too low.
Caves and underground waters hold many unique
and endemic species, particularly of triclads (e.g.,
Kenk, 1987). Some troglobiotic triclads and microturbellarians have recent or remote marine origins (e.g.,
Kawakatsu and Mitchell, 1984a; Kolasa, 1977b). An
extensive study of European underground triclads
(Gourbault, 1972) revealed a suit of distinct distributional, reproductive, and physiological characteristics.
These often include specialized underground microhabitat selection, greater frequency of sexual reproduction,
and slower metabolism as compared to epigean (or
surface-dwelling) triclads.
In contrast, a number of interstitial and underground water microturbellarians, for example Bothrioplana semperi, Limnoruanis romanae, and Stenostomum pegephilum, occur on more than one continent.
Distributions of most genera appear to be worldwide
among microturbellarians, but not triclads, many genera of which are known exclusively from either North
160
Jurek Kolasa
America, Eurasia, South America, or Australia (Ball,
1974).
The majority of triclads from North America are
described from the eastern and southern United States
and Canada (Kenk, 1972). Four triclad species inhabit
Alaskan waters and 10 live in Tennessee (Chandler and
Darlington, 1986). Several species of triclads with recent marine origins occur in freshwater caves of
Mexico (Kawakatsu and Mitchell, 1984b; Benazzi and
Giannini, 1971).
Boreo-alpine distribution of many cold-water species
has been tied to glaciations in Europe and similar patterns also seem to occur in North America (Hampton,
1988). At lower latitudes, similarities between South
American and North American microturbellian composition (cf. Marcus, 1945) suggest considerable interchanges of species.
The ecological distribution of both microturbellarians and triclads has been studied more intensively in
Europe, but most of the results are directly relevant to
North American fauna. The distribution of microturbellarians in lakes has been investigated by Strayer
(1985), Chodorowski (1959), Kolasa (1977a, 1979),
Rixen (1968), Schwank (1976), and Young (1970,
1973a). These studies show that microturbellarians differentiate among substrate types and depth zones. The
flatworm assemblages of sand and gravel bottoms are
distinctly different from assemblages in habitats characterized by bottoms of silt, coarse organic matter, or
vegetation. Plant successional zones also differ in their
species assemblages. Although the littoral zone has the
greatest density and diversity of flatworms, there is apparently no limit to how deep some species may be
found as long as oxygen is available. For example,
Otomesostoma auditivum lives at a depth of 15 – 45 m
in Echo Lake in the Sierra Nevada range, but it occurs
at a depth of 145 m in some European alpine lakes
(Hyman, 1955).
Ecological differentiation of microturbellarians in
running waters is more pronounced than in lakes.
European studies showed distinct assemblages of species
associated with springs, headwaters, and lower stretches
of streams and rivers (epirhithron, metarhithron, hyporhithron, and potamon) (Schwank, 1981; Kolasa,
1983). These regularities in zonation of flatworm assemblages are consistent with patterns observed among
other lotic taxa. These patterns, however, still need to be
related to newer ecological approaches to running waters, i.e., to the river continuum concept (Vannote et al.,
1980). Among those classical zones, epirhithron shows
the greatest species richness. No comparable studies
have been published on North American microturbellarians. Zonation of triclads in streams was studied in a
greater detail, including experimentation (Reynoldson,
1982). Competition, as well as thermal requirements
were implicated as important factors separating several
species of flatworms along a water course and in standing waters (Claussen and Walters, 1982; Pattee, 1980;
Reynoldson, 1982).
Both microturbellarians and triclads live in habitats where the physical conditions require additional
adaptations in comparison to an average pond or
stream environment. Wet mosses and leaves on the water edge, water containers at the base of bromeliad
leaves, the high Arctic, ice melt ponds at elevations
over 4000 m in Central Asia, and hot springs are habitats for one or more species of flatworms. Representatives of freshwater families are also commonly found in
humid terrestrial habitats.
C. Behavioral Ecology
Laboratory studies reveal that turbellarians have a
limited ability to learn simple tasks such as choosing
white over dark branches of a maze (McConnell,
1967). Observations of prey handling (personal observation), however, indicate that their learning ability
may be much greater for tasks that might be advantageous under natural conditions, such as prey or predator recognition.
Possible behavior modification or behavior determination associated with symbiosis is a potentially rewarding research area. Typhloplana viridata, a common spring pond species, is unique in being positively
phototactic. This behavior may be associated with the
presence of symbiotic algae. It might be of general interest to know control mechanisms of this behavior.
D. Foraging Relationships, Predators, and Parasites
Microturbellarians eat bacteria, algae, protozoa,
and small invertebrates, while triclads feed predominantly on larger invertebrates. Scavenging is common
in both morphological groups. Various Mesostoma
species have been studied in detail and illustrate the diversity of feeding habits and techniques within one
genus. Leaf-shaped M. ehrenbergi usually suspend
themselves in the water column on a mucous filament
attached to the surface film (Göltenboth and
Heitkamp, 1977). In that position, the worm waits for
a cladoceran or an insect larva to come into contact.
Using its front end, M. ehrenbergi traps the prey with
sticky mucus, wraps around it, and proceeds to feed.
According to Dumont and Carels (1987), M. lingua
uses neurotoxins to immobilize its prey in addition to
mucus. Mesostoma can also glide on the surface of underwater objects or swim directly in the water column.
Both activities enhance its chances of encountering
6. Flatworms: Turbellaria and Nemertea
161
prey. Unlike M. ehrenbergi, many other species are
spindle-shaped (e.g., M. arctica). These species hunt
more often on the bottom or among submersed objects.
Although several Mesostoma studied spend more than
50% of their time resting on various objects, they can
catch prey as soon as it comes close enough to create
vibrations in the water (J. Kolasa, personal observation). At that point, Mesostoma can erect the front
portion of its body and actively move it around in
search for contact with the prey. Several species of
Mesostoma, such as M. vernale (Hyman) and M. californicum (Hyman), as well as species of Bothromesostoma have a flat ventral surface with which they cling
to the surface film. These species are more agile and
much faster than other species of Mesostoma and they
prefer hunting at the water surface.
Immobilizing the prey and sucking its body fluids
is a common feeding behavior in microturbellarians
equipped with a strong pharynx rosulatus, or in triclads with a pharynx plicatus. Many other turbellarians, however, swallow the whole victim or bite chunks
of prey. For example, species of Stenostomum and
Macrostomum that feed on algae, protozoa, rotifers,
and other meiofauna usually swallow the prey whole.
Other, such as Stenostomum predatorium and various
Phaenocora species may attack and partly consume
larger turbellarians (Kepner and Carter, 1931),
oligochaetes, or other soft invertebrates. Finally, a few
microturbellarians (e.g., Gyratrix hermaphroditus and
Prorhynchus stagnalis) use their copulatory stylets to
stab prey.
Chemical detection of prey undoubtedly plays an
important role in feeding. Injured invertebrates attract
species of both triclads and microturbellarians under
laboratory conditions. The presence of injured or recently killed organisms allows scavenging by many individuals that played no role in immobilizing the prey.
Turbellarians themselves are subject to predation
and parasitism. Ciliates and flagellates are frequent
parasites of Catenulida and Typhloplanida, and
nematodes have been found in Lecithoepitheliata
(Prorhynchus; J. Kolasa, unpublished). Fish occasionally eat triclads (Davies and Reynoldson, 1971). Microturbellarians (Phaenocora typhlops) may occasionally
be eaten by invertebrates, such as a chironomid
Anatopynia (Young, 1973b) or other Turbellaria (e.g.,
Stenostomum predatorium).
(Peter, 1995). The few studies conducted in Britain indicate that intraspecific competition for food may be
very important for Phaenocora typhlops (Young,
1975), while interspecific competition may play a role
in regulating several planarians (Reynoldson, 1982). In
European Polycelis tenuis and, perhaps, in American
Cura foremanii (Girard), this regulation is expressed
by a negative relationship between the worm body size
and density. In fact, long-term changes in the population size of Polycelis suggest that limit cycles or
cyclical fluctuations of the density due to intraspecific
competition are involved (Reynoldson, 1982, and
references therein).
Densities of microturbellaria vary according to the
habitat, season, and species. No major generalizations
have been made, but some patterns are beginning to
emerge, particularly along seasonal and successional
axes. In small, second- or third-order streams, the
greatest densities of microturbellaria are found in the
lower reaches, i.e., in the metarhithron and hyporhithron, while the highest richness is found in the
epirhithron zone (Kolasa, 1983). For example, a mean
density of 1280 individuals / m2 (of all species) was
recorded in Wappinger Creek, New York, in an artifical
substrate after a week of colonization.
Schwank (1981), using volume instead of surface to
quantify turbellarian abundance in submontane streams
in Germany, found densities of about 80 microturbellarians / L. If one assumes an average sample depth into the
substrate of 5 cm, this translates into over 4000 individuals / m2. His data suggest a corresponding density of triclads of 32 individuals / L and 1600 individuals / m2. In
oligotrophic Mirror Lake, turbellarian densities varied
from 40,000 individuals / m2 in shallow littoral regions
to almost zero in profundal zones; the lakewide mean
was 27,000 individuals / m2 (Strayer, 1985). Other studies reported densities of 800 individuals / m2 of Lake
Michigan (Nalepa and Quigley, 1983), 3500 individuals / m2 (annual mean) in shallow littoral areas of Zbechy
Lake in Poland (Kolasa, 1979), 3100 individuals / m2 in
Lake Paajarvi in Finland (Holopainen and Paasivirta,
1977), to 9500 individuals / m2 in ponds in Germany
(Heitkamp, 1982). These values have to be interpreted
cautiously, however, in the light of seasonal succession
and abundance changes, which ordinarily produce the
greatest richness and abundance in early summer
(Bauchhenss, 1971; Strayer, 1985; Chodorowski, 1960).
E. Population Regulation and Density
F. Functional Role in the Ecosystem
It is not clear whether there is any single mode of
population regulation that applies to flatworms. Asexual reproduction of Dugesia tahitiensis was found to
be density-dependent under laboratory conditions
The functional role of species is typically discussed
in terms of their trophic interactions. All triclads are
predatory. Several invertebrates and vertebrates may
consume triclads and are a significant source of their
162
Jurek Kolasa
mortality. However, triclads constitute a relatively
minor component of the diet of their predators (Davies
and Reynoldson, 1971). Triclads themselves consume
isopods, midges, oligochaetes, caddisflies, mayflies, ostracodes, and cladocerans. The ability to exploit various resources differs among species and may result in
strong competition with a simultaneous expansion of
invading species (e.g., Gee and Young, 1993).
Heitkamp (1982) reports intense predation on
Mesostoma and Rhynchomesostoma by Dalyellia in
temporary pools. He has not determined the frequency
of such predation nor established whether this predation results in population regulation.
High densities of both triclads and microturbellarians suggest that their role in biotic interactions of
benthic communities may be greater than their contribution to the diet of other organisms. Densities of
Mesostoma species (lingua) in excess of 1000 individuals / m2 were observed in more than 30% of the
rice fields studied by Collins and Washino (1979) in
California. In some cases, microturbellaria may regulate population dynamics of zooplankton in ponds, as
demonstrated by Maly et al. (1980). They found that
the feeding rates of Mesostoma ehrenbergi were two
zooplankton per day (estimated from Fig. 1). However,
Schwartz and Hebert (1982) reported that this species
can consume about ten cladocerans per day. Blaustein
(1990) and Blaustein and Dumont (1990) found that
Mesostoma lingua might have major numerical impact
on mosquito larvae, cladocerans, and ostracodes in rice
fields, but little effect on copepods. More important,
perhaps, is the functional role of microturbellaria as
consumers of protozoa, rotifers, and algae, especially
by the usually abundant Stenostomum leucops and
similar forms. Unfortunately, no quantitative data on
this subject are available.
Although there are no explict studies of the energy
flow through lake meiobenthos, the detailed study of the
energy budget of Mirror Lake (Strayer and Likens, 1986)
indicated this flow to be equivalent to that to zooplankton. Furthermore, Strayer (1985) showed that Turbellaria, and Rhynchoscolex in particular, were a substantial
component of meiobenthos in Mirror Lake. This statement might be somewhat misleading because meiobenthos itself is a minor component of the whole budget.
IV. CURRENT AND FUTURE RESEARCH
PROBLEMS
A. Dispersal
Understanding the geographical and ecological distribution of Turbellaria is important because their distribution has evolutionary and ecological implications.
Yet this understanding will suffer seriously unless the
dispersal capabilities of flatworms are assessed in terms
of distance and rates. Hebert and Payne (1985) found
low levels of gene flow in Arctic Mesostoma. Their calculated dispersal rates are inconsistent with both the
unpublished data from Ciborowski’s experiment mentioned earlier and with common observations that even
newly created water bodies have a rich complement of
species. A series of questions may be posed. Is dispersal
different at various latitudes? Do dispersal strategies
and efficiency differ along major taxonomic lines? Are
habitat generalists better dispersers than specialists?
Are species associated with a particular habitat type
(e.g., temporary ponds) better dispersers as opposed to
cave dwellers? If yes, what in their life cycle makes
them so? These questions point out the limits in factual
knowledge and ecological interpretation of the many
aspects of turbellarian dispersal.
B. Rhabdoids
Wrona’s (1986) study demonstrated a lack of
support for hypotheses that rhabdoids play a major
role in contacts with prey or predators in triclads. In
many microturbellarians however, rhabdoids, and
adenal products in particular are channeled toward
the anterior of the body. The front end usually plays
a crucial role in exploration of the environment and
in initiation of attack against prey. This special position is highly suggestive of more active applications
of rhabdoids than the mere production of mucus.
Therefore, the role of rhabdoids warrants further
research.
C. Endosymbiosis
Green algae occur in Phaenocora, Dalyellia, Typhloplana, Castrada, and some other microturbellarians. The nature of the association between the algae
and turbellarians is not fully understood. However, it
could be used as a convenient general model for laboratory manipulation.
D. Community Ecology
Comparative research on the density and richness
of turbellarian assemblages in various water types
might suggest which factors have relatively greater influence in structuring flatworm communities. Quantification and standardization of sampling procedures may
be a major hurdle for comparative ecologists. Different
density estimates obtained in quantification of lake
Turbellaria may be due to minor differences in techniques (e.g., Strayer, 1985).
6. Flatworms: Turbellaria and Nemertea
V. COLLECTING, REARING, AND IDENTIFICATION
TECHNIQUES
A variety of techniques can be used to obtain qualitative and quantitative samples of Turbellaria. These
techniques may involve: scooping with a plankton net
over aquatic vegetation or sediments; collecting bottom
sediments by means of bottom samplers, such as Ekman,
Ponar, or multiple corer (Schwank, 1981; Strayer, 1985);
filtering water taken from wells, springs, hyporheic interstitial, or ponds or puddles using a plankton net (Kolasa
et al., 1987); or band baiting (e.g., Mesostoma, triclads)
into traps using pieces of liver or injured aquatic insects
(Case and Washino, 1979; Kenk, 1972). In streams, a
Surber sampler or similar devices can be applied.
Samples should be cooled, topped with water, and
transported to the laboratory as quickly as possible. In
the laboratory, samples can be transferred to jars or
beakers and left for a couple of hours for the water to
stagnate. When the water clears, animals swim toward
the surface or glide on the glass where they can be
picked up with a pipette for further examination. Alternatively, turbellarians can be directly separated from
the sediments under a stereomicroscope, but this procedure is more time consuming. Turbellaria can also be
removed from mineral substrates having a low organic
matter content by gently heating the sample on a hot
plate so that the surface of the sediment reaches
30 – 32°C (Kolasa, 1983). Other methods of extracting
flatworms from sediments were developed for marine
sands (Martens, 1984; Noldt and Wehrenberg, 1984).
These methods are based on various combinations of
anaesthetizing worms (with MgCl2 or ethanol), stirring
water to separate mineral particles, and sieving to catch
worms. Some of these methods may be adaptable to
freshwater fauna.
Many triclads and microturbellarians can be maintained in the laboratory in small glass containers as
long as the water is regularly changed and appropriate
food provided. Synthetic pond water is best for common species. Temperatures between 17 and 25°C may
be adequate for most species, except for cold-water
forms. Food requirements vary with the species and
variety of easily available aquatic invertebrates must be
tested to ensure success. These may include cladocerans, mosquito larvae (for Mesostoma, Dugesia),
oligochaetes (for Phaenocora), protozoa, or other small
turbellarians (Stenostomum) (e.g., Kolasa, 1987; Yu
and Legner 1976). Often, pieces of larger invertebrates,
such as Asellus, Gammarus, and Tubifex, can be successfully used. McConnell (1967) provided many useful
hints on rearing triclads.
Most turbellarians can be identified by squash
mounts of live animals. Identification is difficult for a
163
novice. Squash mounts are prepared by placing a live individual in a drop of water under a microscope coverslip.
Next, excess water is removed with a fine pipette or a
strip of filter paper until the specimen is immobilized.
Pressure on the specimen can be varied by removing or
adding water. Such mounts, with a little bit of practice,
reveal the arrangement and appearance of internal
organs — information usually necessary for species determination. Whenever further study is desirable, animals
can be fixed and preserved as in the following steps.
1. Anaesthetizing with 7% ethanol, 0.1% chloretone, 1% hydroxylamine hydrochloride, or
slowing their locomotion by placing them in a
small volume of water on ice (this step is
optional);
2. Killing with hot fixatives, such as Stieve’s,
Gilson’s, Bouin’s, room temperature 70%
ethanol, or (optional) killing cooled worms with
glutaraldehyde (see electron microscopic
techniques for details);
3. Rinsing mercury residues of the above fixatives
with 50% ethanol solution of iodine (except
Bouin’s);
4. Storing specimens in 70% ethanol.
Animals so prepared can be sectioned and analyzed
with standard histological methods, including staining
with Mallory’s stain, Delafield’s hematoxylin, and
other common stains.
VI. IDENTIFICATION OF NORTH AMERICAN
GENERA OF FRESHWATER TURBELLARIA
The key provided uses a combination of both phylogenetic and superficial morphological and anatomical
characters. Although the phylogenetically important
characters permit greater confidence, they are often
very difficult to use by an inexperienced researcher. It is
strongly recommended that identification be confirmed
by comparing the specimen at hand with exact taxonomic descriptions of the species available in other
publications. Monographs by Luther (1955, 1960,
1963, for Dalyelliidae, Macrostomidae, Typhloplanidae), Nuttycombe (1956), Nuttycombe and
Waters (1938, for Catenula, Stenostomum), Ferguson
(1939, for Macrostomum), and Gilbert (1938, for
Phaenocora) are particularly useful.
A. Taxonomic Key to Orders and Suborders
of Turbellaria
As some characters may be difficult to determine,
it may be advisable to match the general body plan of
164
Jurek Kolasa
the individual being identified with one of the pictures
in Figure 1A – H. For additional schematics one
should consult a guide to world Turbellaria (Cannon,
1986).
1a.
Simple female gonad; egg entolecithal (Acoela, Macrostomida, Catenulida; freshwater members of the latter two often reproduce
asexually and ovary is absent) (Figs. 1A – C, 12A, B)..........................................................................................................................2
1b.
Heterocellular female gonad, with yolk-producing part separate from the oocyte-producing part (all other Turbellaria) (Figs.
1D – H, 2) ...........................................................................................................................................................................................4
2a(1a).
Mouth opens directly or through a pharynx simplex into the body; no protonephridia; no distinct intestine; gonads without clear
walls (Fig. 1A); rare in freshwater..............................................................................................................................................Acoela
[Only one species (unidentified) collected in North America (from Mirror Lake in New Hampshire; Strayer, 1985).]
2b.
Mouth opens to a pharynx simplex; protonephridia present; epithelial, ciliated intestine present.......................................................3
3a(2b).
Protonephridia with a single, central excretory duct, often two or more zooids; male gonopore, if present, situated on the dorsal
side and in the front of the body; ovary without oviducts and supplementary organs; statocyst present in some species (Figs. 1B,
12A, B) ...................................................................................................................................................Catenulida (see Section VI.B)
3b.
Protonephridia with a pair of main excretory ducts; male gonopore on the ventral side of the posterior part of body; female gonopore usually separate and in front of the male one, more than one zooid in the family Microstomidae only; no statocyst (Figs. 1C,
5A, B) ...............................................................................................................................................Macrostomida (see Section VI.C)
4a(1b).
Single germovitellarium where ova are surrounded by yolk cell epithelium; pharynx variabilis; copulatory organ armed with a sclerotized stylet near the pharynx (Figs. 1D, 2A)..............................................................................Lecithoepitheliata (see Section VI.D)
4b.
Ova not surrounded by yolk epithelium; ovary separate or combined with vitellarium (Figs. 1E – H, 2B, C)......................................5
5a(4b).
Pharynx plicatus or variabilis; testicles and ovaries dispersed; germ cells dispersed or aggregated (Fig. 1H)..................Proseriata, Prolecithophora, and Tricladida (see Section VI.E)
5b.
Pharynx bulbosus (doliiformis or rosulatus, Fig. 1E – G) ....................................................................................................................6
6a(5b).
Pharynx doliiformis, directed forward (Fig. 1E) ...................................................................................Dalyellioida (see Section VI.H)
6b.
Pharynx rosulatus directed ventrally or, exceptionally as in Phaenocora, ventrally and forward ........................................................7
7a(6b).
Without a proboscis (Fig. 1F) ..........................................................................................................Typhloplanoida (see Section VI.F)
7b.
Proboscis present (Fig. 1G) ............................................................................................................Kalyptorhynchia (see Section VI.G)
FIGURE 2
Examples of different types of heterocellular
female gonads. (A) Prorhynchus, Geocentrophora; (B)
Bothrioplana; and (C) Typhloplana, Dalyellia.
6. Flatworms: Turbellaria and Nemertea
165
B. Taxonomic Key to Catenulida
1a.
Brain simple, compact, oval ...............................................................................................................................................................2
1b.
Brain composed of paired frontal and posterior lobes; a group of sensory cells in front of the brain, often arranged pseudometamerically, i.e., in parallel rows .........................................................................................................................family Stenostomidae
5
2a(1a).
Brain situated usually at the base of, but not further than in the middle of prostomium (frontal body section without the intestine);
with ciliated ventral or lateroventral furrow separating prostomium from the rest of body ...........................family Catenulidae
3
2b.
Brain situated in the first 1/3 of prostomium ..............................................................................family Chordariidae
3a(2a).
A ring of strongly ciliated longitudinal grooves developed on the prostomium (Fig. 3A) ........................................................Suomina
3b.
No such grooves present ....................................................................................................................................................................4
4a(3b).
Clearly open intestine extends, approximately, to the middle of the postpharyngeal section of body (Figs. 3B,C, 12A)..........Catenula
4b.
Open intestine extends much further, leaving no more than 1/5 to 1/4 of the postpharyngeal section closed; usually the intestinal
lumen of individual zooids is connected ...........................................................................................................................Dasyhormus
5a(1b).
Adult individuals composed of two or more zooids; gonads often absent...........................................................................................6
5b.
Adult individuals (with gonads) without signs of asexual divisions; with long prostomium, ciliated pits absent (Figs. 3D,F, 12B)........
.....................................................................................................................................................................................Rhynchoscolex
6a(5).
A section of the intestine near the pharynx with a strong ring of muscles (Fig. 4E) ..................................................Myostenostomum
6b.
No such structure, intestine uniformly ciliated (Fig. 4A – D)............................................................................................Stenostomum
FIGURE 3
Some representatives of Catenulida. (A) Suomina turgida; (B) Catenula lemnae;
(C) Catenula leptocephala; and (D) and (E) Rhynchoscolex simplex. Length of species varies
between 0.35 and 5.00 mm. B, Brain; f, furrow; i, intestine; m, mouth; n, protonephridial
duct; o, ovary; ph, pharynx; and st, statocyst.
Chordarium
166
Jurek Kolasa
FIGURE 4
Some representatives of Catenulida. (A) Stenostomum leucops; (B) S. beauchampi; (C)
S. glandulosum; (D) S. brevipharyngium; and (E) Myostenostomum tauricum. B, Brain; cp, ciliated
sensory pits; g, muscular ring of intestine or gizzard; i, intestine; lo, light-refracting organs; m,
mouth; n, protonephridial duct; pg, pharyngeal glands; ph, pharynx; and tc, tactile cilia.
C. Taxonomic Key to Macrostomida
1a.
Ciliated pits present; intestine extends in front of mouth; numerous zooids separated by division planes (Fig. 5B,C) family Microstomidae .............................................................................................................................................................................Microstomum
1b.
No ciliated pits; intestine entirely behind the pharynx; no signs of asexual division (Fig. 5A, D, E) ..................family Macrostomidae
Macrostomum
D. Taxonomic Key to Lecithoepitheliata
1a.
Stylet of the copulatory organ straight; no eyes ................................................................................................................Prorhynchus
1b.
Stylet of the copulatory organ curved; eyes present except in some cave forms ...........................................................Geocentrophora
E. Taxonomic Key to Prolecithophora, Proseriata, and Tricladida
1a.
Mouth situated in front and directed forward ...................................................................................................................Hydrolimax
1b.
Mouth situated elsewhere and directed backward or ventrally ...........................................................................................................2
2a(1b).
Mouth situated near the middle and directed ventrally; statocysts present....................................................................Otomesostoma
2b.
Mouth directed backward; no statocyst .............................................................................................................................................3
6. Flatworms: Turbellaria and Nemertea
167
FIGURE 5 Representatives of Macrostomida and copulatory organs of various families. (A) Macrostomum
tuba; (B) Microstomum lineare; (C) copulatory organ of M. lineare; (D) stylet of Macrostomum gilberti; (E)
stylet of an unidentified Macrostomum from New Haven, CT; (F) sclerotized copulatory apparatus of Microdalyellia rossi (Dalyelliidae); (G) sclerotized copulatory apparatus of M. tennesseensis; and (H) copulatory
organ of Opistomum pallidum containing a spiny reversible cirrus (Typhloplanidae). Ci, Cirrus; co, copulatory organ; cp, ciliated pits; e, eye; i, intestine; m, mouth; o, ovary; ov, egg; ph, pharynx; ps, prostatic secretions; s, sperm; t, testes; and vs, seminal vesicle.
3a(2b).
Intestine split into two branches behind the pharynx (Figs. 6 – 7) ........................................................................................Tricladida1
3b.
Intestine not composed of three branches (Fig. 8A)
4a(3a).
Internal pharyngeal muscles in two distinct layers (Planariidae) (Fig. 6) .............................................................................................5
4b.
Internal pharyngeal muscles in one layer of mixed longitudinal and circular fibers (Dendrocoelidae) (Fig. 6)...................................12
5a(4a).
Oviducts, separate or united, open into end part of the bursa stalk ....................................................................................................6
5b.
Oviducts united, the common oviduct opens in the genital atrium (Fig. 6) .........................................................................................7
6a(5a).
Testes extend to the level of pharynx ............................................................................................................................................Cura
6b.
Testes extend to the posterior end of body...............................................................................................................................Dugesia
7a(5b).
Numerous eyes arranged in a band around the anterior end of body (Fig. 7G) .......................................................................Polycelis
7b.
Eyes absent or one pair only; if numerous, then not along the head margin .......................................................................................8
8a(7b).
Adenodactyl present (Fig. 6)....................................................................................................................................................Planaria
8b.
Adenodactyl absent ............................................................................................................................................................................9
9a(8b).
Anterior end with an adhesive organ ................................................................................................................................................10
9b.
Anterior end without an adhesive organ...........................................................................................................................................11
10a(9a).
Body elongated, flat, with a well-developed postpharyngeal section .................................................................................Sphalloplana
1
Substantial parts of the key to Tricladida follow Kenk (1972). Separation of Hymanella from Phagocata is based on Ball et al. (1981).
168
Jurek Kolasa
FIGURE 6
Schematic representation of the major anatomic features used in taxonomy of
Tricladida. Ad, Adenodactyl; b, bursa; bd, bursal duct; ca, common genital atrium; im, internal muscle layer of the pharynx (two possible types shown); m, mouth; ma, male atrium; odc,
common oviduct; ph, pharynx; pb, penis bulb; pp, penis papilla; sd, sperm duct; v, vagina;
and vs, seminal vesicle.
10b.
Body turtle-shaped, with reduced postpharyngeal section .........................................................................................................Kenkia
11a(9b).
Prepharyngeal, fused testes in male phase; large atrium lined by papillose epithelium in female phase; head round and broad; the
anterior ramus of the intestine does not reach beyond eyes .................................................................................................Hymanella
11b.
Other set of characters .........................................................................................................................................................Phagocata
12a(4b).
Anterior end of the body with a deeply invaginated adhesive organ .................................................................................Macrocotyla
12b.
Adhesive organ absent, or an adhesive disk present..........................................................................................................................13
FIGURE 7
Some common representatives of Tricladida. (A) Hymanella
retenuova; (B) Phagocata velata; (C) Dugesia tigrina; (D) D. dorotocephala;
(E) D. polychroa; (F) Procotyla fluviatilis; and (G) Polycelis coronata (A and
B modified from Ball et al., 1981).
6. Flatworms: Turbellaria and Nemertea
169
FIGURE 8 Representatives of (A) Proseriata (Bothrioplana semperi); (B) Typhloplanoida
(Mesosomta craci); and (C – F) Kalyptohynchia; (C) Gyratrix hermaphroditus and (D) its
stylet; (E) Opisthocystis goettei and (F) its copulatory organs. Ar, Adenal rhabdoids; b,
brain; co, copulatory organ; cp, ciliated pits; dr, dermal rhabdoids; e, eye; ev, excretory vesicle; i, intestine; n, protonephridial duct; o, ovary; pg, pharyngeal glands; ph, pharynx; pr,
proboscis; s, stylet; t, testes; vg, granular vesicle; vs, seminal vesicle; and y, yolk glands.
13a(12b).
Penis papilla present .........................................................................................................................................................................14
13b.
Penis papilla absent, bulb large........................................................................................................................................Rectocephala
14a(13a).
Penis bulb rounded, containing a seminal vesicle........................................................................................................Dendrocoelopsis
14b.
Penis bulb elongated, contains a prostatic vesicle ..................................................................................................................Procotyla
F. Taxonomic Key to Typhloplanoida
The basic plan of the freshwater typhloplanoid reproductive system is given in Fig. 9.
1a.
Yolk glands paired, situated on both sides of the intestine..................................................................................................................2
1b.
Yolk glands developed as single strand of cells above the intestine (Fig. 10E, F)...............................................................Limnoruanis
2a(1a).
Pharynx developed as a typical, round or slightly elongated pharynx rosulatus and oriented ventrally or somewhat forward ...........3
2b.
Pharynx elongated or cylindrical, oriented backward; copulatory organ with a spiny cirrus (Fig. 5H) ..............................Opistomum
3a(2a).
Testes under yolk glands.....................................................................................................................................................................4
3b.
Testes above yolk glands ..................................................................................................................................................................11
4a(3a).
Protonephridial ducts open separately on the body surface (Protoplanellinae) (as in Fig. 10C) ..........................................................5
4b.
Protonephridial ducts combined with mouth or gonopore (as in Fig. 10A) ........................................................................................7
5a(4a).
Front extremity with a central depression and set slightly apart by two lateral, ciliated pits; pharynx in the first third of body (Fig.
10C) Prorhynchella
5b.
Front extremity not set apart, rounded; pharynx in the second to third portion of body ....................................................................6
6a(5b).
Reproductive complex with a bursa copulatrix; ductus ejaculatorius usually surrounded by gelatinous matrix (Fig. 11E, F; cf.,
Fig. 9) ...............................................................................................................................................................................Krumbachia
6b.
Bursa copulatrix absent ...................................................................................................................................................Amphibolella
7a(4b).
Protonephridial ducts joined and combined with mouth; no retractable proboscis (Typhloplaninae) .................................................8
170
Jurek Kolasa
FIGURE 9 A schematic representation of the main reproductive organs and structures of freshwater
Typhloplanoida. In some species, additional structures may be present or the relative arrangement among
the organs may be slightly different.
7b.
Protonephridial ducts open into the gonopore; retractable front extremity (Fig. 10D) ..........................................Rhynchomesostoma
8a(7a).
With a bursa copulatrix and an atrium copulatorium (Fig. 11D)............................................................................................Castrada
8b.
One of the above organs missing ........................................................................................................................................................9
9a(8b).
With eyes (Figs. 10A, 12C) ..........................................................................................................................................Strongylostoma
9b.
No eyes ............................................................................................................................................................................................10
10a(9b).
Atrium copulatorium absent; bursa copulatrix present..................................................................................................Typhloplanella
10b.
Both bursa copulatrix and atrium copulatorium absent; always with green algae; pharynx in the first half of body; copulatory organ
pear-shaped.......................................................................................................................................................................Typhloplana
11a(3b).
Pharynx developed as a typical, round pharynx rosulatus ................................................................................................................13
11b.
Pharynx barrel-shaped, directed clearly forward, and situated near the front extremity (Fig. 11B)...................................................12
12a(11b).
Ductus copulatorius absent; inferior genital atrium inconspicuous (Fig. 11B) ....................................................................Phaenocora
12b.
With ductus copulatorius; exceptionally large inferior atrium; occurs in sulfuric springs ........................................Pseudophaenocora
13a(11a).
An elongated supplementary organ, the ascus, present in the proximity of the gonopore; or with a well-defined gland complex in its
place (Ascophorinae) (cf. Fig. 9) .......................................................................................................................................................14
13b.
Ascus absent.....................................................................................................................................................................................16
14a(13a).
Pharynx in the middle of the first half of body ....................................................................................................................Protoascus
14b.
Pharynx approximately in the middle of body or slightly behind .....................................................................................................15
15a(14b).
Gonopore near the middle of body ......................................................................................................................................Ascophora
6. Flatworms: Turbellaria and Nemertea
FIGURE 10
Some representatives of Typhloplanoida. (A) Strongylostoma simplex; (B) copulatory organ with
testes of S. simplex; (C) Prorhynchella minuta; (D) a juvenile of Rhynchomesostoma rostratum; (E) Limnoruanis romanae; and (F) sclerotized part of the copulatory organ, partly reverted (upper picture), and the copulatory organ with the gonopore and glands (lower picture) in L. romanae (C modified from Rusebush, 1939). Ar,
Adenal rhabdoids; e, eye; g, glands; i, intestine; n, protonephridial duct; o, ovary; pg, pharyngeal glands; ph,
pharynx; s, stylet,; sph, sphincter; t, testes; vg, granular vesicle; vs, seminal vesicle; and y, yolk glands.
FIGURE 11 Representatives of the Dalyellioida (Castrella) and Typhloplanida (others); (A) Castrella
pinguis; (B) Phaenocora sp. (from Churchill, Manitoba); (C) Mesostoma vernale (the habitus of this
species is also characteristic of various Bothromestoma); (D) Castrada virginiana; (E) Krumbachia
hiemalis; and (F) copulatory complex of Krumbachia hiemalis. ar, Adenal rhabdoids; bc, copulatory
bursa; bs, blind sac of the atrium; co, copulatory organ; de, ductus ejaculatorius; gp, gonopore; i,
intestine; m, matrix; n, protonephridial duct; o, ovary; ov, egg capsule; ph, pharynx; r, seminal receptacle; s, stylet; t, testes; vs, seminal vesicle; and y, yolk glands.
171
172
Jurek Kolasa
FIGURE 12
Microphotographs of live Turbellaria and Nemertea. (A) Several partly framed individuals of
Catenula sp. showing chains of zooids; (B) frontal portion of body of Rhynchoscolex sp.; (C) Strongylostoma
sp., a representative of Typhloplanoida; (D) squash preparation of the portions of the reproductive system of
Mesostoma craci, with the copulatory organ in the middle; and (E) a frontal portion of body of a nemertean
Prostoma sp. B, Brain; BC, bursa copulatrix; CD, common duct; ED, ejaculatory duct; prostatic granulations
are visible in its proximity; I, intestine; M, mouth; N, protonephridium; O, ovary; PH, pharynx; R, rhynchocoel; S, sensory pit; ST, statocyst; T, testicle; Y, yolk glands; I, first zooid; and II, second zooid.
6. Flatworms: Turbellaria and Nemertea
173
15b.
Gonopore in the third posterior part of body ...............................................................................................................Dochmiotrema
16a(13b).
Protonephridial ducts open separately on the body surface; eyes often present..................................................................Olisthanella
16b.
Protonephridial ducts open into mouth ............................................................................................................................................17
17a(16b).
With a ventral pit and a canal connecting the bursa copulatrix to the ovovitellin duct; often flat on the ventral side and strongly
convex on the dorsal ................................................................................................................................................Bothromesostoma
17b.
Without the above characters (Figs. 11C, 8B, 12D) ............................................................................................................Mesostoma
G. Taxonomic Key to Kalyptorhynchia and Similar Forms
1a.
Excretory vesicle absent; protonephridial ducts open directly at the surface of body; single ovary (Fig. 8C).......................................2
1b.
Protonephridial ducts open into an excretory vesicle in the rear end of body; paired ovary (Fig. 8E, F).........................Opisthocystis2
2a(1a).
Copulatory organ with an almost straight stylet (Fig. 8C, D) .................................................................................................Gyratrix
2b.
Copulatory organ with a curved stylet.............................................................................................................Microkalyptorhynchus3
H. Taxonomic Key to Dalyellioida
1a.
With a paired ovary; copulatory organ with a straight, funnel-like tube..............................................................................Pilgramilla
1b.
With a single ovary; copulatory organ with a more of less complex sclerotized structure (e.g., Figs. 5F, G, 11A) ...............................2
2a(1b).
Sclerotized structure directly attached to the rest of the copulatory organ ..........................................................................................3
2b.
Sclerotized structure in a separate pocket adjacent to the bulb of the copulatory organ (Fig. 11A).........................................Castrella
3a(2a).
Sclerotized structure typically with two handles and two lateral branches carrying spines, auxiliary parts may be present (Fig. 5F, G)
...........................................................................................................................................................................................................4
3b.
Sclerotized structure different; usually in the form of a spiny ring ......................................................................................Gieysztoria
4a(3a).
One to several eggs in the parenchyme; normally, older individuals green due to symbiotic zoochlorellae..............................Dalyellia
4b.
Never more than one egg; no zoochlorellae ...................................................................................................................Microdalyellia
NEMERTEA
VII. GENERAL CHARACTERISTICS, EXTERNAL
AND INTERNAL ANATOMICAL FEATURES
Nemerteans are coelomate (Turbeville and Ruppert, 1985) and unsegmented worms possessing an
eversible muscular proboscis resting in the rhynchocoel, a walled, longitudinal dorsal cavity connected
with the mouth (Fig. 13 and 14, see also Figure 12E in
Section VI.F.4 Unlike the Turbellaria, Nemertea have an
anus as well as a closed-blood circulatory system. A
monolayered, ciliated epidermis and several layers of
muscles enclose the body. The excretory system is protonephridial. Sensory structures include eyes, statocysts
2
An unidentified kalyptorhynchid occurring in a tributary of the
Hudson River (Kolasa et al., 1987) also meets this criterion, although it may belong to a family other than that containing
Opisthocystis. Klattia is a synonym of Opisthocystis (Karling T.G.
1956. Ark. Zool. 9:187 – 279).
3
Taxonomic placement within Kalyptorhynchia is provisional. Name
(absent in freshwater nemerteans), and cerebral,
frontal, and lateral neural organs. The reproductive
system is much simpler than in Turbellaria. Freshwater
nemerteans are hermaphroditic and often protandric.
Spermatozoa and ova are produced in numerous gonads situated in diverticulae of the intestine. As a rule,
fertilization is external and self-fertilization can also
occur. Development is direct. Most information provided in this section on nemerteans is based on an
excellent review of freshwater and terrestrial nemerteans by Moore and Gibson (1985). That review
will be an invaluable source of data and references for
anyone working on the ecology and biology of freshwater nemerteans.
discussed by W.E. Hazen (1953. Morphology, taxonomy, and
distribution of Michigan rhabdocoeles. Ph.D. Thesis, Univ. of
Michigan, Ann Arbor. 122 pp.).
4
Until 1985, Nemertea were considered to be acoelomate. With the
new finding (Turbeville and Ruppert, 1985), they can no longer be
associated with other flatworms. Our presentation here is more for
convenience than systematic reasons.
174
Jurek Kolasa
FIGURE 14
Sections through the anterior extremity of three North
American nemerteans. (A) Prostoma eilhardi; (B) P. graecense; (C) P.
canadiensis (after Gibson and Moore, 1976, 1978). bw, Body wall
musculature; cg, cephalic gland lobular region; dc, dorsal cerebral
commissure; ep, epidermis; fo, frontal organ; id, improvised ducts
from cephalic gland lobules; oe, oesophagus; pr, proboscis; rc, rhynchocoel; rd, rhynchodaeum; rl, rhychodaeal longitudinal muscle
fibers; st, stomach; and vc, ventral cerebral commisure.
VIII. ECOLOGY
A. Life History and General Ecology
FIGURE 13
A semi-schematic view of Prostoma sp. b, Brain; e,
eye; fo, frontal organ; g, gonad; i, intestine; ov, egg; s, stylet.
Twelve species of freshwater nemerteans are classified into two, possibly polyphyletic families and six
genera (Moore and Gibson, 1985). In the family Heteronemertea, three monotypic genera are recognized:
Planolineus (Beauchamp, 1928), Siolineus (du BoisReymond Marcus, 1948), and Apartronemertes
(Wilfert and Gibson, 1974). In the family Hoplonemertea, the genus Prostoma (Duges, 1828) contains
most of the species. Two other genera, Campbellonemertes (Moore and Gibson, 1972) and Potamonemertes (Moore and Gibson, 1973) are known from islands in the southern Pacific, Campbell Island and
South Island, New Zealand, respectively.
Most nemerteans are marine, although a small minority are terrestrial or freshwater. The freshwater
forms are benthic, predatory worms 10 – 40 mm in
length and of pink coloration. Prostoma feed readily
on oligochaete worms and, occasionally, on crustaceans, nematodes, turbellarians, midge larvae, and
other small invertebrates (McDermott and Roe, 1985).
Feeding activity is most intense at night and the sticky
proboscis captures prey. Little is known about habitat
selection by freshwater nemerteans. North American
Prostoma species have been found in both streams and
lakes, including Lake Huron. In lakes, Prostoma
species appear to be associated with filamentous algae.
Even though some nemerteans have strong regenerative capabilities, reproduction is exclusively sexual,
with both male and female organs present at the same
time. Since only limited quantitative studies have been
conducted on the biology of freshwater nemerteans in
6. Flatworms: Turbellaria and Nemertea
the field, mechanisms of population regulation remain
unknown. Clearly, Prostoma can reproduce rapidly
with up to 210 eggs per reproductive episode throughout the year as long as the temperature is above 10°C
(Young and Gibson, 1975). A study of one population
in England suggests that Prostoma population density
increases during the cold months of the year and declines during the summer, with smaller individuals
recorded during the winter (Gibson and Young, 1976).
It is not clear, however, if this reproduction pattern is
general or due to cattle trampling of the shore vegetation where the study was conducted.
Distribution of nemerteans in North America is
not well known. (Strayer, 1985) reported Prostoma
rubrum from Mirror Lakes, New Hampshire. However, P. rubrum is no longer a valid species (Gibson and
Moore, 1976), and this observation has to be treated
as Prostoma sp. Prostoma canadiensis is known from
one site in Lake Huron (Gibson and Moore, 1978),
where is occurs to a depth of 20 m. Prostoma species
are probably much more common in freshwaters than
the infrequent records from eastern North American
might indicate. A Prostoma species (P. rubra) was also
reported in vegetation of slow coastal streams,
marshes, and estuarine pools in St. Vincent, Caribbean
(Harrison and Rankin, 1976). So far, no Prostoma
have been found in temporary water bodies.
Three established species of nemerteans are known
from North America (Fig. 14); a fourth, P.
asensoriatum, is a questionable species. Distribution of
these species in North America is summarized in Figure
15, but they are probably present in most standing and
running waters. With the exception of the dubious
P. asensoriatum, all three remaining species have been
175
recorded in other parts of the world, including South
America, Europe, Africa, and Australia. Zoologists do
not know whether the worldwide pattern of distribution is natural or, instead, a result of recent humanmediated dispersal. Analysis of genetic diversity of various populations might shed some light on this question.
B. Physiological Adaptations
Osmoregulation is an ecologically important function in nemerteans as in all other freshwater invertebrates with permeable body walls. It is controlled by
the cerebral organs and involves several organs and enzymatic systems associated with blood vessels (Moore
and Gibson, 1985). Well-developed nephridia may play
a role in osmoregulation as well as in the removal of
nitrogenous wastes (but the latter has not yet been
demonstrated). Gases, particularly oxygen, are exchanged across the surface of the body. The role of
blood vessels in oxygen transport is unclear.
Nemerteans, like turbellarians, release copious mucus whose various functions probably include defense,
locomotion, physiological barrier, and encystment.
C. Behavioral Ecology
There are no studies devoted exclusively to the behavior of freshwater nemerteans. Nevertheless, some
observations on feeding behavior, escape reactions, and
locomotion are available. Adult nemerteans can only
crawl, while small juveniles also swim. Forward locomotion of Prostoma may involve ciliary movement
alone, ciliary movement in combination with muscular
waves, sinusoidal curves when the animal pushes
through the vegetation, or peristalsis when the animal
moves backward. The proboscis provides the fastest
movement forward. This long organ is rapidly everted
and attached to substrate, and the body is then pulled
toward the point of attachment.
Typically, prey are captured by a sticky proboscis
and then swallowed whole by the widely distended
mouth (McDermott and Roe, 1985). The mechanisms
of prey detection are poorly understood. In terrestrial
nemerteans, these mechanisms are unusual in that they
involve ambushing strategy and reliance on mechanical
stimuli.
D. Functional Role in the Ecosystem
FIGURE 15
Known distribution of Prostoma species in North
America (after Gibson, 1985; modified and updated). C, P. canadiensis; E, P. eilhardi; A, P. asensoriatum; G, P. graecense; and U, unidentified Prostoma.
Necessarily, the functional importance of nemerteans in the ecosystem can only be indirectly inferred from their trophic position and relative densities.
I have found densities of Prostoma species in Wappinger Creek, a tributary of the Hudson River, to vary
176
Jurek Kolasa
between 50 and 590 individuals / m2, which constitutes
a small fraction of all predatory invertebrates identified
at the study site. Similarly, low densities of P. canadiensis (up to 140 individuals / m2) were observed on gravelly and muddy substrates in Lake Huron.
IX. CURRENT AND FUTURE
RESEARCH PROBLEMS
In view of the limited knowledge of nemertean
ecology, almost any area of research will provide valuable information. New information on phenology, population dynamics, and habitat selectivity would permit
a better evaluation of the role of nemerteans in freshwater communities. Specific studies in the biology of
nemerteans appear particularly promising in the areas
of reproductive biology and evolutionary ecology. The
ecological role of femaleness later in the life cycle, the
production of resting eggs, and the environmental versus genetic cues of life stages all may offer exciting and
general models for evolutionary ecology.
X. COLLECTION, CULTURING, AND PRESERVATION
Collection of nemerteans is similar to collection of
any soft-bodied, benthic invertebrates such as turbellari-
ans or oligochaetes living in the substrate or on aquatic
plants. Sieving and vigorous washing may damage specimens. Placing samples in beakers or jars and topping
with water allows nemerteans to crawl toward the edges
of water where they can be collected with a pipette.
Freshwater nemerteans are relatively easy to culture. They may be maintained and will reproduce in
aquaria or even in petri dishes in clean, oxygenated water and on a diet of live aquatic invertebrates. Chopped
tubificid oligochaetes provide a good diet.
Worms can be preserved in 80% ethanol. Before
preservation, however, worms should be narcotized using 7% ethanol or chloretone until their movement
ceases. According to Moore and Gibson (1985), gin
can be used in the field if pure ethanol is unavailable,
but only as a last resort!
XI. TAXONOMIC KEY TO SPECIES OF
FRESHWATER NEMERTEA IN NORTH AMERICA
It should be noted that Gibson and Moore (1976,
1978) provided keys to freshwater nemerteans of the
world. Those keys are more suitable if you encounter a
species that has not yet been recorded in North America. The present key is an adaptation of the keys by
Gibson and Moore (1976, 1978) pertinent to established North American Prostoma.
1a.
Cephalic glands open via frontal organs and by improvised ducts (Fig. 14C); esophagus distinct but unciliated; rhynchodaeum with
isolated longitudinal muscle strands; proboscis with twelve nerves. P. canadiensis (Gibson and Moore, 1978)
1b.
Cephalic glands open only via frontal organ, no improvised ducts .....................................................................................................2
2a(1b).
With a distinctive ciliated esophagus; cephalic glands reach back to the brain (Fig. 14B); rhynchodaeum with a well-developed layer
of longitudinal muscle ..............................................................................................................................P. graecense (Bohmig, 1892)
2b.
Different combination of characters; indistinct and unciliated esophagus (Fig. 14A); rhynchodaeum without specifically associated
layer of longitudinal muscle fibers; proboscis with 9 – 10 nerves..........................................................P. eilhardi (Montgomery, 1894)
ACKNOWLEDGMENTS
I am very grateful to L.R.G. Cannon, J. Moore, R.
Gibson, A.P. Mead, R. Kent, S. Schwartz, D. Strayer,
and S. Tyler for helpful comments on the manuscript,
additional references, and permission to reproduce figures, glossary entries, and fragments of keys.
LITERATURE CITED
Ball, I. R. 1974. A contribution to the phylogeny and biogeography
of the freshwater triclads (Platyhelminthes: Turbellaria), in:
Riser, N.W., Morse, M.P. Eds., Biology of the Turbellaria. McGraw-Hill, New York, pp. 339 – 401.
Bauchhenss, J. 1971. Die Kleinturbellarien Frankens. Ein Beitrag zur
Systematik und Ökologie der Turbellaria excl. Tricladida in
Süddeutschland. Internationale Revue des gesamten Hydrobiologie 56:609 – 666.
Bedini, C., Ferrero, E., Lanfranchi, A. 1973. Fine structure of the
eyes in two species of Dalyelliidae (Turbellaria Rhabdocoela).
Monitore Zoologico Italiano 7:51 – 70.
Benazzi, M., Giannini, E. 1971. Cura azteca, nuova specie di planaria del
Messico. Atti della Accademia Nazionale dei Lincei 50:477–481.
Blaustein, L. 1990. Evidence for predatory flatworms as organizers of
zooplankton and mosquito community structure in rice fields.
Hydrobiologia 199:179 – 191.
Blaustein, L., Dumont, H. J. 1990. Typhloplanid flatworms (Mesostoma and related genera): mechanisms of predation and evidence that they structure aquatic invertebrate communities. Hydrobiologia 198:61 – 77.
Boddington, M. J., Mettrick, D. F. 1977. A laboratory study of the
population dynamics and productivity of Dugesia polychroa
(Turbellaria: Tricladida). Ecology 58:109 – 118.
Calow, P., Read, D. A. 1986. Ontogenetic patterns and phylogenetic
trends in freshwater flatworms (Tricladida); constraint or selection? Hydrobiologia 132:263 – 272.
Cannon, L. R. G. 1986. Turbellaria of the world. A guide to families
and genera, Queensland Museum, Brisbane, QLd., Australia.
6. Flatworms: Turbellaria and Nemertea
Case, T. J., Washino, R. K. 1979. Flatworm control of mosquito larvae in rice fields. Science 206:1412 – 1414.
Chandler, C. M., Darlington, J. T. 1986. Further field studies on
freshwater planarians of Tennessee (Turbellaria: Tricladida): III.
Western Tennessee. Journal of Freshwater Ecology 3:493 – 501.
Chodorowski, A. 1959. Ecological differentiation of turbellarians in
Harsz Lake. Polskie Archiwum Hydrobiologii 6:33 – 73.
Chodorowski, A. 1960. Vertical stratification of Turbellaria species
in some littoral habitats of Harsz Lake. Polskie Archiwum Hydrobiologii 8:153 – 163.
Claussen, D. L., Walters, L. M. 1982. Thermal acclimation in the
freshwater Planarians, Dugesia tigrina and D. dorotocephala.
Hydrobiologia 94:231 – 236.
Collins, F. H., Washino, R. K. 1979. Factors affecting the density of
Culextarsalis and Anopheles freeborni in northern California
rice fields. Proceedings of California Mosquito Control Association 46:97 – 98.
Davies, R. W., Reynoldson, T. B. 1971. The incidence and intensity
of predation on lake-dwelling triclads in the field. Journal of
Animal Ecology 40:191 – 214.
Dumont, H. J., Carels, I. 1987. Flatworm predator (Mesostoma cf.
lingua) releases a toxin to catch planktonic prey (Daphnia
magna). Limnology and Oceanography 32:699 – 702.
Dumont, H. J., Schorreels, S. 1990. A laboratory study of the feeding
of Mesostoma lingua (Schmidt) (Turbellaria: Neorhabdocoela)
on Daphnia magna Straus at four different temperatures. Hydrobiologia 198:79 – 89.
Ehlers, U. 1986. Comments on a phylogenetic system of the Platyhelminthes. Hydrobiologia 132:1 – 12.
Ferguson, F. F. 1939. A monograph of the genus Macrostomum O.
Schmidt 1848. Part II. Zoologischer Anzeiger 126:131 – 144.
Fiore, L., Ioalé, P. 1973. Regulation of the production of subitaneous
and dormant eggs in the turbellarian Mesostoma ehrenbergii
(Focke). Monitore Zoologico Italiano 7:203 – 224.
Gee, H., Young, J. O. 1993. The food niches of the invasive Dugesia
tigrina (Girard) and indigeneous Polycelis tenuis Ijima and P.
nigra (Müller) (Turbellaria; Tricladida). Hydrobiologia 254:
99 – 106.
Gibson, R., Moore, J. 1976. Freshwater nemerteans. Zoological
Journal of the Linnean Society 58:177 – 218.
Gibson, R., Moore, J. 1978. Freshwater nemerteans: new records of
Prostoma and a description of Prostoma canadiensis sp. nov.
Zoologischer Anzeiger 201:77 – 85.
Gibson, R., Young, J. O. 1976. Ecological observations on a population of the freshwater hoplonemertean Prostoma jenningsi Gibson and Young 1971. Archiv für Hydrobiologie 78:42 – 50.
Gilbert, C. M. 1938. Two new North American Rhabdocoeles —
Phaenocora falciodenticulata nov. spec. and Phaenocora kepneri adenticulata nov. subspec. Zoologischer Anzeiger
122:208 – 223.
Göltenboth, F., Heitkamp, U. 1977. Mesostoma ehrenbergi (Focke,
1836) Plattwürmer (Strudelwürmer). Biologie, mikroskopische
Anatomie und Cytogenetik, in: Siewing, R., Ed., Grosses Zoologisches Praktikum Heft 6a:pp. 1 – 60.
Gourbault, N. 1972. Recherches sur les triclades paludicoles hypogés.
Memories du Muséum. National d’Histoire Naturelle 73:1 – 249.
Hampton, A. M. 1988. Altitudinal range and habitat of triclads in
streams of the Lake Tahoe basin. American Midland Naturalist
120:302 – 312.
Harrison, A. D., Rankin, J. J. 1976. Hydrobiological studies of Eastern Lesser Antillean Islands. II. St. Vincent: freshwater fauna —
its distribution, tropical river zonation and biogeography.
Archiv für Hydrobiologie Suppl 50:275 – 311.
Hebert, P. D. N., Payne, W. 1985. Genetic variation in populations of
the hermaphroditic flatworm Mesostoma lingua (Turbellaria,
177
Rhabdocoela). Biological Bulletin (Woods Hole, MA)
169:143 – 151.
Heitkamp, U. 1979a. Abundanzdynamik und reproduktionsphasen
limnophiler tricladen (Turbellaria) aus stehenden kleingewassern. Hydrobiologia 65:49 – 57.
Heitkamp, U. 1979b. Die Respirationsrate neorhabdocoeler Turbellarien mit unterschiedlicher Temperaturvalenz. Archiv für Hydrobiologie 87:95 – 111.
Heitkamp, U. 1982. Untersuchungen zur Biologie, Ökologie und Systematik limnischer Turbellarien periodischer und perennierender
Kleingewässer Südniedersachsens. Archiv für Hydrobiologie,
Supplement 64:65 – 188.
Holmquist, Ch. 1967. Turbellaria of northern Alaska and northwestern Canada. Internationale Revue des gesamten Hydrobiologie
52:123 – 139.
Holopainen, I. J., Paasivirta, L. 1977. Abundance and biomass of the
meiozoobenthos in the oligotrophic and mesohumic lake Paajarvi, southern Finland. Annales Zoologici Fennici 14:124 – 134.
Hyman, L. H. 1951. The invertebrates: Platyhelminthes and Rhynchocoela, the acoelomate Bilateralia. McGraw-Hill, New York.
Hyman, L. H. 1955. Descriptions and records of freshwater Turbellaria from the United States. American Museum Novitates
1714:1 – 36.
Jennings, J. B. 1977. Patterns of nutrition in free-living and symbiotic
Turbellaria and their implications for the evolution of parasitism in the phylum Platyhelmithes. Acta Zoologica Fennica
154:63 – 79.
Jennings, J. B. 1985. Feeding and digestion in the aberrant planarian
Bdellasimilis barwicki (Turbellaria:Tricladidae:Procerodidae):
an ectosymbiote of freshwater turtles in Queensland and New
South Wales. Australian Journal of Zoology 33:317 – 327.
Kawakatsu, M., Mitchell, R. W. 1984a. A list of retrobursal triclads
inhabiting freshwaters and aquatic probursal triclads regarded
as marine relicts, with corrective remarks on our 1984 paper
published in Zoological Science, Tokyo. Occasional Publications, Biological Laboratory of Fuji Women’s College, Sapporo
11:1 – 8.
Kawakatsu, M., Mitchell, R. W. 1984b. Redescription of Dugesia
azteca (Benazzi et Giannini, 1971) based upon the material
collected from the type locality in Mexico, with corrective
remarks. Bulletin of the National Science Museum, Ser. A
10:37 – 50.
Kenk, R. 1972. Freshwater planarians (Turbellaria) of North America, in: EPA Biota of Freshwater Ecosystems, Identification manual 1:1 – 81.
Kenk, R. 1987. Freshwater triclads (Turbellaria) of North America.
XVI. More on subterranean species of Phagocata of the Eastern
United States. Proceedings of the Biological Society of Washington 100:664 – 673.
Kepner, W. A., Carter, J. S. 1931. Ten well-defined new species of
Stenostomum. Zoologischer Anzeiger 98:108 – 123.
Kolasa, J. 1977a. Bottom fauna of the heated Konin Lakes. Turbellaria and Nemertini. Monografie Fauny Polski 7:29 – 48.
Kolasa, J. 1977b. Remarks on the marine-originated Turbellaria in
the freshwater fauna. Acta Zoologica Fennica 1954:81 – 87.
Kolasa, J. 1979. Ecological and faunistical characteristics of Turbellaria in the eutrophic Lake Zbechy. Acta Hydrobiologica
21:435 – 459.
Kolasa, J. 1983. Formation of the turbellarian fauna in a submontane stream. Acta Zoologica Cracoviensia 26:57 – 107.
Kolasa, J. 1987. Population growth in some Mesostoma species
(Turbellaria), predatory on mosquitoes. Freshwater Biology
18:205 – 212.
Kolasa, J., Strayer, D., Bannon-O’Donnell, E. 1987. Microturbellarians from interstitial waters, streams, and springs in southeastern
178
Jurek Kolasa
New York. Journal of the North American Benthological Society
6:125 – 132.
Lanfranchi, A., Papi, F. 1978. Turbellaria (excl. Tricladida), in: Illies,
J. Ed., Limnofauna Europaea, G. Fisher, Stuttgart.
Luther, A. 1955. Die Dalyelliiden. Acta Zoologica Fennica
87:1 – 337.
Luther, A. 1960. Die Turbellarien Ostfennoskandiens. I. Acoela,
Catenulida, Macrostomida, Lecithoepitheliata, Prolecithophora,
und Proseriata. Fauna Fennica 7:1 – 155.
Luther, A. 1963. Die Turbellarien Ostfennoskandiens IV. Neorhabdocoela 2. Typhloplanoida: Typhloplanidae, Solenopharyngidae
und Carcharodopharyngidae. Fauna Fennica 16:1 – 163.
Maly, E. J., Schoenholtz, S., Arts, M. T. 1980. The influence of flatworm predation on zooplankton inhabiting small ponds. Hydrobiologia 76:233 – 240.
Marcus, E. 1945. Sobre microturbellarios do Brasil. Comunicaciones
Zoologicas del Museo de Historia Natural de Montevideo
25:1 – 97.
Marcus, E. 1951. Contributions to the natural history of Brazilian
Turbellaria. Comunicaciones Zoologicas del Museo de Historia
Natural de Montevideo 63:1 – 25.
Martens, P. M. 1984. Comparison of three different extraction methods for Turbellaria. Marine Ecology Progress Series 14:
229 – 234.
McConnell, J. V. 1967. A manual of psychological experimentation
on planarians. A special publication of the Worm Runner’s Digest. J. V. McConnell, Ann Arbor, MI.
McDermott, J. J., Roe, P. 1985. Food, feeding behavior and feeding
ecology of nemerteans. American Zoologist 25:113 – 125.
Mead, A. P., Kolasa, J. 1984. New records of freshwater Microturbellaria from Nigeria, West Africa. Zoologischer Anzeiger
212:257 – 271.
Moore, J., Gibson, R. 1985. The evolution and comparative physiology of terrestrial and freshwater nemerteans. Biological Reviews
60:257 – 312.
Nalepa, T. F., Quigley, M. A. 1983. Abundance and biomass of
meiobenthos in nearshore Lake Michigan with comparisons to
macrozoobenthos. Journal of the Great Lakes Research
9:523 – 529.
Noldt, U., Wehrenberg, C. 1984. Quantitative extraction of living
Platyhelminthes from marine sands. Marine Ecology Progress
Series 20:193 – 201.
Nuttycombe, J. W. 1956. The Catenula of the eastern United States.
American Midland Naturalist 55:419 – 433.
Nuttycombe, J. W., Waters, A. J. 1938. The American species of the
genus Stenostomum. Proceedings of the American Philosophical
Society 79:213 – 301.
Pattee, E. 1980. Coefficients thermiques et ecologie de quelques
planaires d’eau douce. VII: leur zonation naturelle. Annales de
Limnologie 16:21 – 41.
Peter, R. 1995. Regenerative and reproductive capacities of the fissiparous planarian Dugesia tahitiensis. Hydrobiologia 305:261.
Reynoldson, T. B. 1982. The population biology of Turbellaria with
special reference to the freshwater triclads of the British Isles.
Advances in Ecological Research 13:235 – 365.
Rieger, R. M. 1986. Asexual reproduction and the turbellarian archetype. Hydrobiologia 132:35 – 45.
Rixen, J.-U. 1968. Beitrag zur Kenntnis der Turbellarienfauna des
Bodenseegebietes. Archiv für Hydrobiologie 64:335 – 365.
Ruiz-Trillo, I., Riutort, M., Littlewood, D. T. J., Herniou, E. A.,
Baguñà, J. 1999. Acoel flatworms: earliest extant bilateralian
metazoans, not members of Platyhelminthes. Science
283:1919 – 1923.
Sayre, R. M., Powers, E. M. 1966. A predacious soil turbellarian that
feeds on free-living and plant-parasitic nematodes. Nematologica 12:619 – 629.
Schwank, P. 1976. Quantitative Untersuchungen an litoralen Turbellarien des Bodensees. Jh. Ges. Naturkde. Württemberg 131:
163 – 181.
Schwank, P. 1981. Turbellarien, Oligochaeten and Archianneliden
des Breitenbachs und anderer oberchessischer Mittelgebirgsbäche. II. Die Systematik und Autökologie der einzelnen Arten.
Archiv für Hydrobiologie, Supplement 62:1 – 85.
Schwartz, S. S., Hebert, P. D. N. 1982. A laboratory study of the
feeding behaviour of the rhabdocoel Mesostoma ehrenbergii on
pond Cladocera. Canadian Journal of Zoology 60:1305 – 1307.
Schwartz, S. S., Hebert, P. D. N. 1986. Prey preference and utilization by Mesostoma lingua (Turbellaria, Rhabdocoela) at a low
arctic site. Hydrobiologia 135:251 – 257.
Strayer, D. 1985. The benthic micrometazoans of Mirror Lake,
New Hampshire. Archiv für Hydrobiologie, Supplement 72:
287 – 426.
Strayer, D., Likens, G. E. 1986. An energy budget for the zoobenthos
of Mirror Lake, New Hampshire. Ecology 67:303 – 313.
Turbeville, J. M., Ruppert, E. E. 1985. Comparative ultrastructure
and evolution of Nemertines. American Zoologist 25:53 – 71.
Tyler, S., Burt, M. D. B. 1988. Lensing by a mitochondrial derivative
in the eye of Urastoma cyprinae (Turbellaria, Prolecithophora).
Fortschritte der Zoologie 36:229 – 234.
Vannote, R. L., Minshall, G. W., Cummins, K. W., Sedell, J. R.,
Cushing, C. E. 1980. The river continuum concept. Canadian
Journal of Fisheries and Aquatic Sciences. 37:130 – 137.
Wrona, F. 1986. Distribution, abundance, and size of rhabdoids in
Dugesia polychroa (Turbellaria: Tricladida). Hydrobiologia
132:287 – 293.
Young, J. O. 1970. British and Irish freshwater Microturbellaria: historical records, new records and a key for their identification.
Archiv für Hydrobiologie 67:210 – 241.
Young, J. O. 1973a. The occurence of microturbellaria in some
British lakes. Archiv für Hydrobiologie 72(2):207 – 208.
Young, J. O. 1973b. The prey and predators of Phaenocora typhlops
(Vejdovsky) (Turbellaria: Neorhabdocoela) living in a small
pond. Journal of Animal Ecology 42:637 – 643.
Young, J. O. 1975. The population dynamics of Phaenocora typhlops (Vejdovsky) (Turbellaria: Neorhabdocoela) living in a
pond. Journal of Animal Ecology 44:251 – 262.
Young, J. O., Eaton, J. W. 1975. Studies on the symbiosis of Phaenocora typhlops (Vejdovsky) (Turbellaria; Neorhabdocoela) and
Chlorella vulgaris var. vulgaris, Fott & Novakova (Chloroccales). II. An experimental investigation into the survival value
of the relationship to host and symbiont. Archiv für Hydrobiologie 75:225 – 239.
Young, J. O., Gibson, R. 1975. Some ecological studies on two populations of the freshwater hoplonemertean Prostoma eilhardi
(Montgomery, 1894) from Kenya. Verhandlungender internationale Vereinigung für. Limnologie 19:2803 – 2810.
Yu, H.-S., Legner, E. F. 1976. Regulation of aquatic diptera by
planaria. Entomophaga 21:3 – 12.
6. Flatworms: Turbellaria and Nemertea
APPENDIX 6.1 List of North American Species of Microturbellaria
The arrangement of orders and families follows
with some modifications from that of Lanfranchi and
Papi (1978).
Catenulida
Family Catenulidae
Catenula confusa
Catenula lemnae
Catenula leptocephala
Catenula sekerai
Catenula virginia
Suomina turgida
Family Chordariidae
Chordarium europaeum
Family Stenostomidae
Myostenostomum tauricum
Rhynchoscolex platypus
Rhynchoscolex simplex
Rhynchoscolex sp. 1
Rhynchoscolex sp. 2
Stenostomum anatirostrum
Stenostomum anops
Stenostomum arevaloi
Stenostomum beauchampi
Stenostomum brevipharyngium
Stenostomum ciliatum
Stenostomum cryptops
Stenostomum glandulosum
Stenostomum grande
Stenostomum kepneri
Stenostomum leucops
Stenostomum mandibulatum
Stenostomum membranosum
Stenostomum occultum
Stenostomum pegephilum
Stenostomum predatorium
Stenostomum pseudoacetabulum
Stenostomum simplex
Stenostomum temporaneum
Stenostomum tuberculosum
Stenostomum unicolor
Stenostomum uronephrium
Stenostomum ventronephrium
Stenostomum virginianum (probably
synonymous with S. unicolor)
Macrostomida
Family Macrostomidae
Macrostomum collistylum
Macrostomum curvistylum
Macrostomum gilberti
Macrostomum glochostylum
Macrostomum lewisi
Macrostomum norfolkense
Macrostomum ontarioense
Macrostomum orthostylum
Macrostomum phillipsi
Macrostomum reynoldsi
Macrostomum riedeli
Macrostomum ruebushi
Macrostomum sensitivum
Macrostomum sillimani
Macrostomum sp. 1, indeterm.
Macrostomum sp. 2, indeterm.
Macrostomum sp. 3, indeterm.
Macrostomum stirewalti
Macrostomum tenneseense
Macrostomum tuba
Macrostomum lineare
Macrostomum virginianum
Family Microstomidae
Microstomum lineare
Lecithoepitheliata
Family Prorhynchidae
Geocentrophora cavernicola
Geocentrophora sphyrocephala
Prorhynchus stagnalis
Proseriata
Family Plagiostomidae
Hydrolimax grisea
Prolecithophora
Family Bothrioplanidae
Bothrioplana semperi
Family Otomesostomidae
Otomesostoma auditivum
Dalyellioida
Family Provorticidae
Genus indet. species indet
Pilgramilla virginiensis
Family Dalyelliidae
Castrella pinguis
Castrella graffi
Dalyellia viridis
Microdalyella abursalis
Microdalyella circobursalis
Microdalyella fairchildi
Microdalyella gilesi
Microdalyella rheesi
Microdalyella rochesteriana
Microdalyella rossi
Microdalyella ruebushi
Microdalyella sillimani
Microdalyella tennesseensis
Microdalyella virginiana
Typhloplanida
Family Typhloplanidae
Amphibolella spinulosa
179
180
Jurek Kolasa
Ascophora elegantissima
Bothromesostoma personatum
Castrada hofmanni
Castrada lutheri
Castrada virginiana
Castrada sp. indet
Krumbachia cf. hiemalis
Krumbachia minuta
Krumbachia virginiana
Limnoruanis romanae
Mesostoma arctica
Mesostoma californicum
Mesostoma columbianum
Mesostoma craci
Mesostoma curvipenis
Mesostoma ehrenbergii
Mesostoma macroprostatum
Mesostoma platygastricum
Mesostoma vernale
Mesostoma virginianum
Olisthanella truncula
Opistomum pallidum
Phaenocora agassizi
Phaenocora falciodenticulata
Phaenocora highlandense
Phaenocora kepneri
Phaenocora lutheri
Phaenocora virginiana
Prorhynchella minuta
Protoascus wisconsinensis
Pseudophaenocora sulflophila
Rhynchomesostoma rostratum
Strongylostoma cf., elongatum
Strongylostoma radiatum
Typhloplanella halleziana
Kalyptorhynchia
Family Polycystidae
Gyratrix hermaphroditus
Opistocystis goettei
Family undetermined
Microrhynchus virginianus
7
GASTROTRICHA
David Strayer
William D. Hummon
Institute of Ecosystem Studies
Millbrook, New York 12545
Department of Biological Sciences
Ohio University
Athens, Ohio 45701
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Organ System Function
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Collecting, Rearing, and Preparation for
Identification
V. Taxonomic Key
Literature Cited
I. INTRODUCTION
The gastrotrichs are among the most abundant and
poorly known of the freshwater invertebrates. Gastrotrichs are nearly ubiquitous in the benthos and periphyton of freshwater habitats, with densities typically
in the range of 100,000 – 1,000,000 individuals/m2.
Nonetheless, the remarkable life cycle of freshwater
gastrotrichs was worked out only recently, and we
know almost nothing about how the distribution and
abundance of these animals are controlled in nature.
The impact of freshwater gastrotrichs on their food resources and on freshwater ecosystems has not yet been
investigated. Twelve genera and fewer than 100 species
of freshwater gastrotrichs are now known from North
America. Because the North American gastrotrich
fauna has received so little study, these numbers understate the real diversity of the fauna.
Formerly placed in the Aschelminthes or Nemathelminthes, gastrotrichs usually are now considered to
constitute a phylum of their own. The evolutionary
placement of gastrotrichs is unclear; some authors
(e.g., Lorenzen, 1985; Wallace et al., 1996) believe
that they are most closely related to nematodes, kinorhynchs, loriciferans, nematomorphs, and priaulids,
while others (e.g., Conway Morris, 1993) think that
gastrotrichs are more closely related to rotifers and
acanthocephalans. Important general references on
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
gastrotrichs include Remane (1935 – 1936), Hyman
(1951), Voigt (1958), d’Hondt (1971a), Hummon
(1982), Ruppert (1988), Schwank (1990), and
Kisielewski (1991, 1998). The phylum contains two
orders: the Macrodasyida, which consists almost
entirely of marine species, and the Chaetonotida,
which contains marine, freshwater, and semiterrestrial
species. Unless noted otherwise, the information included in this chapter refers to freshwater members of
the Chaetonotida.
Macrodasyids usually are distinguished from
chaetonotids by the presence of pharyngeal pores and
more numerous adhesive tubules (Fig. 1). Macrodasyids are common in marine and estuarine sands, but
are barely represented in inland freshwaters. Two
species of freshwater gastrotrichs have been placed in
the Macrodasyida.
Ruttner-Kolisko (1955) described an aberrant
gastrotrich, Marinellina flagellata (Fig. 1A), from the
hyporheic zone of an Austrian river. Unfortunately,
she was able to find only two specimens, both of
them apparently immature. Because Marinellina has a
pair of anterior lateral structures that Ruttner-Kolisko
interpreted as adhesive tubules, she placed this species
in the Macrodasyida. Remane (1961) rejected the
assignment of Marinellina to the macrodasyids and
placed it instead in the chaetonotid family Dichaeturidae. Kisielewski (1987) reaffirmed Ruttner-Kolisko’s
181
182
David Strayer and William D. Hummon
A. External Morphology
Gastrotrichs are colorless animals, spindle- or tenpin-shaped, and ventrally flattened (Fig. 2). Freshwater
gastrotrichs are 50 – 800 m long. Conspicuous external features include a more or less distinct head, which
bears sensory cilia, and a cuticle which, in most
species, is ornamented with spines or scales of various
shapes. In the most common freshwater family, the
Chaetonotidae (as well as in the rare Dichaeturidae and
Proichthydiidae), the posterior end of the body is
formed into a furca, which contains distal adhesive
tubes that allow the animal to attach itself tenaciously
to surfaces. In other families, these structures are
absent, but the posterior end of the body may bear
long spines or sensory bristles. The ventral side of the
animal bears longitudinal rows or patches of cilia,
which provide the forward-gliding locomotion of the
animal.
B. Organ System Function
FIGURE 1
Freshwater macrodasyid gastrotrichs: (A) Marinellina
flagellata, and (B) Redudasys fornerise. AT, adhesive tube; and PP,
pharyngeal pore. From Ruttner-Kolisko (1955) and Kisielewski
(1987).
original placement of the species. An animal similar to
Marinellina was just discovered in the hyporheic zone
of another Austrian stream (Schmid-Araya and Schmid,
1995); but until it is studied critically, the systematic
placement of these enigmatic Austrian gastrotrichs will
remain unclear.
Kisielewski (1987) discovered an undoubted macrodasyid, Redudasys fornerise (Fig. 1B), from the psammon of a Brazilian reservoir. Additional freshwater
macrodasyids probably will be found when appropriate
habitats (psammon, hyporheic zone) are explored. The
distribution, biology, and evolutionary relationships of
any such species will be of great interest.
II. Anatomy and Physiology
The following brief account of gastrotrich anatomy
is summarized chiefly from Remane (1935 – 1936),
Hyman (1951), Hummon (1982), and Boaden (1985),
which should be consulted for greater detail.
The digestive system begins with a subterminal
mouth, which may be surrounded by a ring of short
bristles. Between the mouth and the pharynx lies a
cuticular buccal capsule, which is often somewhat
protrusible. The muscular pharynx is similar to the nematode pharynx, with a triradiate, Y-shaped lumen.
Often, there are anterior and posterior swellings, but
these lack the valves characteristic of the pharyngeal
bulbs of nematodes. Posterior to the pharynx is an undifferentiated gut, which empties into the anus.
The paired reproductive organs lie lateral to the
gut in the posterior half of the body. In young animals,
large, developing, parthenogenetic eggs are present.
As there are no oviducts, the egg is released through
a rupture in the ventral body wall. Older animals
become hermaphrodites (see below) and bear both
sperm sacs and developing sexual (i.e., meiotic) eggs
lateral to the gut. An unpaired organ of unknown
function, the X-organ, lies posterior to the gonads near
the anus.
The brain is bilobed, straddling the pharynx. Sensory organs include long cilia and bristles, which presumably are tactile. Balsamo (1980) demonstrated that
Lepidodermella squamata is photosensitive, although it
does not appear to have distinct multicellular photoreceptors. Gray and Johnson (1970) found evidence of a
tactile chemical sense in a marine macrodasyid; it
seems probable that freshwater chaetonotids possess
similar chemosensory abilities.
The excretory system consists of a pair of protonephridia (Brandenburg, 1962) in the anterior mid-
7. Gastrotricha
183
FIGURE 2
Schematic illustration of a typical chaetonotid gastrotrich showing (A) dorsal view, (B) ventral
view, (C) posterior end of a hermaphrodite, showing sexual organs, and (D) egg. AT, adhesive tubes; AN, anus;
BR, brain; E, egg; F, furca; G, gut; M, mouth; PH, pharynx; PN, protonephridium; SC, sensory cilia; SS, sperm
sac; and X, X-organ. Modified from Remane (1935 – 1936), Voigt (1958), and Kisielewska (1981).
body, which empty through pores on the ventral body
surface. There is no circulatory or respiratory system,
per se.
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
Gastrotrichs are widely distributed in freshwaters
in surface sediments and among vegetation. Some
species of the Neogosseidae and Dasydytidae are good
swimmers and are occasionally reported from the
plankton of shallow, weedy lakes (e.g., Hutchinson,
1967; Green, 1986; Kisielewski, 1991). However, none
of the gastrotrichs has become as truly planktonic as
the daphnid cladocerans or ploimate rotifers.
Gastrotrichs have been found in a wide range of
freshwater and semiterrestrial habitats. They are abundant in most lakes, ponds, and wetlands (Tables I and
II). Kisielewski (1981, 1986a) showed that gastrotrich
density and species richness are positively correlated
with the productivity of the habitat, and several workers have found that gastrotrich density is highest in
highly organic sediments (e.g., Strayer, 1985). Apparently, most gastrotrichs live very near the sediment
184
TABLE I
David Strayer and William D. Hummon
Number of Species of Gastrotrichs Found in Some Freshwater Habitats.
Habitat
Total
Chaetonotus
European lakesa
18
12
Mirror Lake, NH
Ponds, Polandb
20 – 32
21
8 – 20
10
Peat bogs, Polandc
Bog pools, Polandd
Phragmites mats,
Romania
Ponds, Brazile
Rivers, Brazile
27
24
28
38
22
Other
Chaetonotidae
Dasydytidae
Neogosseidae
Source
5
1
0
12
7
0
3
0
1
16
12
16
9
8
7
2
4
5
0
0
0
Preobrajenskaja, 1926; Balsamo,
1981, 1990, Bertolani and
Balsamo, 1989
Strayer, 1985
Nesteruk, 1986; Kisielewski,
1986a
Kisielewski, 1981
Kisielewska, 1982
Rudescu, 1968
14
5
12
12
10
4
2
1
Kisielewski, 1991
Kisieleswki, 1991
a
Mean of four lakes.
Mean of seven ponds.
c
Mean of four bogs.
d
Mean of two pools.
e
Sum of several collecting sites.
b
surface in lakes (Fig. 3). Gastrotrichs also are abundant
in unpolluted streams, where they inhabit sand bars,
sometimes in great numbers (Hummon et al., 1978;
Hummon, 1987). Apparently, they are scarce in
groundwaters other than the hyporheic zone (RenaudMornant, 1986). Their rarity in underground waters is
surprising, because many marine gastrotrichs are interstitial in habit (d’Hondt, 1971a; Renaud-Mornant,
1986), and because the small size and bacterial diet of
gastrotrichs would seem to preadapt them to the
groundwater habitat.
Gastrotrichs are among the few animals commonly
found in anaerobic environments (Moore, 1939; Cole,
1955; Strayer, 1985), remaining abundant even during
TABLE II
extended periods (i.e., months) of anoxia. The physiological basis of the anaerobiosis of freshwater gastrotrichs has not yet been studied. It seems likely that
some freshwater gastrotrichs possess a sulfide detoxification mechanism similar to that demonstrated for marine
gastrotrichs (Powell et al., 1979, 1980) and freshwater
nematodes (Nuss, 1984; Nuss and Trimkowski, 1984)
to deal with the elevated concentrations of H 2S that
often accompany extended anoxia.
Little is known of the factors that control the
distribution of individual species of gastrotrichs in
freshwater. Arguing largely from analogy with marine
work (d’Hondt, 1971a), we might expect factors of primary importance to include the granulometry, stability,
Density and Biomass of Gastrotrichs in Some Freshwater Habitats.
Site
Density (No. / m2)
Biomass (mg DM / m2)
Source
Lake Bikcze, Poland
Lake Brzeziczno, Poland
Lake Piaseczno, Poland
Mirror Lake, NH
Lake Suviana, Italy
Lake Erie, OH
Three small ponds, Poland
Mississippi River, MN
1,160,000a
920,000 a
910,000 c
130,000 d
57,000 e
50,000 f
1,600,000 – 2,600,000
130,000 – 230,000g
23a,b
30a,b
23b,c
1d
—
—
25 – 78b
—
Nesteruk, 1996
Nesteruk, 1996
Nesteruk, 1996
Strayer, 1985
Madoni, 1989
Evans, 1982
Nesteruk, 1996
Hummon, 1987, and unpublished
a
Littoral zone.
Converted from wet mass by multiplying by 0.15.
c
Mean of three stations.
d
Lakewide mean.
e
Mean of two deepwater stations.
f
Beaches.
g
Sand bars.
b
7. Gastrotricha
FIGURE 3 Vertical distribution of gastrotrichs within the sediments
of Lake Brzeziczno, Poland, (dashed line) and Mirror Lake, New
Hampshire, (solid line) as a function of depth from the sediment surface. From data of Strayer (1985) and Nesteruk (1991).
packing, and organic content of the sediment, the
amount of dissolved oxygen, and the density and composition of communities of microbes and predators.
Also, culture work suggests that the inorganic chemistry of the water and the presence of anthropogenic
contaminants can exert a strong influence on gastrotrich populations (Hummon, 1974; Faucon and
Hummon, 1976; Hummon and Hummon, 1979).
Many genera of freshwater gastrotrichs are known
to have intercontinental or cosmopolitan distributions.
Exceptions include several genera so far known only
from Brazil (the macrodasyid Redudasys, the
chaetonotids Arenotus and Undula, and the dasydytid
Ornamentula); Dichaetura and Marinellina, rare
species known only from Europe, and the proichthydiids Proichthydium and Proichthyioides, which have
been found only at their type localities in Argentina
and Japan, respectively. The geographic distribution of
individual species is not well known, because of the
primitive state of the species-level taxonomy of freshwater gastrotrichs and the paucity of field studies
throughout most of the world. Even in North America,
only Arkansas (Davis, 1937), Illinois (Robbins, 1965,
1973; Horlick, 1975), northern Indiana (Pfaltzgraff,
1967), Michigan (Brunson, 1947, 1948, 1950), and
eastern Ohio (Craft, 1993) have received even cursory
185
surveys of freshwater gastrotrichs. Thus, the 76 species
that Schwank (1990) reported from North America
probably are a small fraction of North America’s freshwater gastrotrich fauna. Some species have been reported to occur over broad ranges, including in some
cases more than one continent (e.g., Kisielewski, 1991).
Until further studies are made, such reported intercontinental distributions should be regarded with caution
(cf. Frey, 1982; Todaro et al., 1996), especially because
“conspecifics” collected from different continents often
exhibit marked morphological differences from one another (e.g., Robbins, 1973, Emberton, 1981).
Chaetonotus typically dominates gastrotrich faunas everywhere in freshwater, both in terms of numbers
of species and numbers of individuals (Table I). Other
genera of the Chaetonotidae are common in all kinds
of freshwaters, but are usually less abundant than
Chaetonotus. The Dasydytidae and Neogosseidae are
less widespread than the Chaetonotidae, usually living
in weedy, productive waters, where they may, however,
become numerically abundant (e.g., Blinn and Green,
1986; Kisielewski, 1981, 1986a; Nesteruk, 1986).
Dasydytids and neogosseids also are more abundant
and speciose in the tropics than in temperate waters
(Table I; Kisielewski 1991). The very rarely seen
Dichaeturidae and Proichthydiidae have been collected
from cisterns, underground waters, tree holes, and
among moss (Remane, 1935 – 1936, Ruttner-Kolisko,
1955; Sudzuki, 1971).
B. Reproduction and Life History
Until recently, populations of freshwater gastrotrichs were thought to consist entirely of parthenogenetic females (e.g., Hyman; 1951; Pennak, 1978).
However, more detailed recent studies have dispelled
this notion, and have revealed a remarkable life cycle
among freshwater gastrotrichs (Fig. 4) (Weiss and Levy,
1979; Hummon, 1984a – c, 1986; Levy, 1984).
Newly hatched gastrotrichs are relatively large (approximately two-thirds the length of adults — Brunson,
1949) and already contain developing parthenogenetic
eggs. These eggs develop rapidly under favorable
conditions. At 20°C, the first egg may be laid within a
day after the mother hatches. Typically, a total of four
parthenogenetic eggs is laid over a 4-day period. Apparently, parthenogenesis is apomictic, so that offspring are genetically identical to their mother.
There are two kinds of parthenogenetic eggs. The
more common kind, tachyblastic eggs, develop immediately and hatch quickly (within a day of being laid, at
20°C). Occasionally, the final parthenogenetic egg laid
by a female is not a tachyblastic egg, but a resting, or
opsiblastic, egg. Opsiblastic eggs are thick-shelled, a
186
David Strayer and William D. Hummon
FIGURE 4
Schematic diagram of the proposed generalized life cycle for freshwater chaetonotid gastrotrichs, based predominately on study of Lepidodermella squamata. Dashed lines show hypothetical
events which have not yet been demonstrated. From Levy (1984), after ideas presented by Levy and Weiss
(1980), with permission of the authors. OPSI, opsiblastic egg; and TACHY, tachyblastic egg.
little larger than tachyblastic eggs, and are very resistant to freezing and drying (Brunson, 1949). The factors that induce the production of opsiblastic eggs are
not well known, although such eggs are often produced
by animals in crowded cultures. Opsiblastic eggs are almost always the final egg produced by an animal, even
if the total number of eggs is fewer than four.
Following the production of parthenogenetic eggs,
animals develop into hermaphrodites (Fig. 2C) (Hummon and Hummon, 1983; 1988). During this time,
sperm and meiotic sexual eggs are produced, and the
X-body grows. These changes occur slowly, over the
period of a week after the last parthenogenetic egg is
laid. No one has yet observed sperm transfer or fertilization, although Levy and Weiss (Levy and Weiss,
1980; Levy, 1984) reported finding a third kind of egg
(the “plaque-bearing egg”) in cultures of Lepidodermella squamata. They suggested that plaque-bearing
eggs may be the product of sexual reproduction. Because the sperms are few in number (32 – 64 per animal) and nonmotile, fertilization probably is internal.
Animals reared in isolation do not appear to produce
fertilized sexual eggs, so cross-fertilization probably is
the rule. The absence of ducts associated with the male
or female reproductive system makes it difficult to
suggest a mechanism of sperm transfer (Hummon
1986, described one bizarre possibility). Probably the
enigmatic X-organ is involved. Much remains to be
learned about the postparthenogenetic sexual phase
and its importance in nature.
The life cycle just described is unique among invertebrates and offers considerable ecological flexibility to
gastrotrich populations. The initial parthenogenetic
phase allows for explosive population growth under
favorable conditions: workers have commonly reported
growth rates (r) of 0.1 – 0.5 per day in laboratory
cultures (e.g., Hummon, 1974; Faucon and Hummon,
1976; Hummon and Hummon, 1979; Hummon, 1986).
Production of parthenogenetic resting eggs (opsiblastic
eggs) buffers the population against unfavorable conditions and presumably allows for dispersal among habitats. Finally, the subsequent sexual phase introduces genetic recombination. Sexual reproduction is most likely
to occur in populations in which rates of mortality are
low enough to allow some gastrotrichs to reach the age
required for sexual development.
C. Ecological Interactions
Gastrotrichs feed on bacteria, algae, protozoans,
detritus, and small inorganic particles. Bacteria probably are of primary importance. Bennett (1979) demonstrated that Lepidodermella squamata readily digested
bacteria and found that this gastrotrich would not
7. Gastrotricha
survive in laboratory cultures in the absence of bacteria. He reported that L. squamata could digest the
green alga Chlorella as well, but suggested that algae
were of secondary importance in gastrotrich diets.
Gray and Johnson (1970) showed that the marine
macrodasyid Turbanella hyalina could choose among
various strains of natural bacteria, apparently on the
basis of a tactile chemical sense. Thus, the quality as
well as quantity of bacterial populations was important
to the gastrotrichs. Freshwater gastrotrichs are most
likely capable of similar fine discrimination among bacteria and other prey.
Reported predators of gastrotrichs include heliozoan and sarcodine amoebae, cnidarians, and tanypodine midges (Brunson, 1949; Bovee and Cordell, 1971;
Moore, 1979), but many other benthic predators presumably feed on gastrotrichs. Nothing is known about
the importance of predation in regulating populations
of freshwater gastrotrichs or about the quantitative importance of gastrotrichs as a food item for various
predators.
We have no direct information on what regulates
gastrotrich populations in nature. Gastrotrichs are
capable of enormous population growth (10 – 50%
per day) in laboratory cultures. If potential growth
rates are anywhere near this high in nature, as seems
likely in some circumstances, then there must be an
equally high counterbalancing mortality, perhaps
from predation. There are no detailed studies of the
population dynamics of freshwater gastrotrichs in nature. The few quantitative studies of the seasonal dynamics of freshwater gastrotrich populations (Fig. 5;
see also Nesteruk, 1986) have shown that population
densities are usually (but not always) lowest during
the winter. We do not know what drives these sea-
FIGURE 5 Seasonal trends in gastrotrich abundance. Left: density
(number / cm3) of two species of dasydytids in the surface sediments
of a boggy pool (“Complex B”) in Poland: Setopus dubius (䊉) and
Dasydytes ornatus (䉱). From data of Kisielewska (1982). Right: density (number / cm2) of all gastrotrichs (䊉), an unidentified species
(probably of Heterolepidoderma) (䊏), and Lepidodermella triloba
(䉱) on the gyttja sediments of Mirror Lake, New Hampshire (original). Plotted points are means s.e. (n 16).
187
sonal dynamics, but seasonal changes in water temperature, food supply, and predation pressures are obvious possibilities.
Gastrotrichs, along with nematodes and rotifers,
are among the most abundant animals in the freshwater benthos, often having densities on the order of
10 – 100 individuals/cm2 ( 100,000 – 1,000,000 individuals/m2) (Table II). However, because there have
been no direct measurements of the roles of gastrotrichs in freshwater ecosystems, their importance
can only be guessed at. There is only a single, tentative
estimate of gastrotrich metabolism in freshwater:
Strayer (1985) estimated secondary production and
respiration of gastrotrichs in Mirror Lake, New
Hampshire, each to be 50 – 100 mg dry mass per
square meter per year, which is less than 1% of total
production or respiration of the zoobenthic community. This estimate suggests that gastrotrich metabolism, and processes such as nutrient regeneration
that are correlated with metabolism, are of minor importance in freshwater ecosystems. It would be imprudent, however, to dismiss gastrotrichs as quantitatively
unimportant to ecosystem functioning without actually
measuring gastrotrich activities under defined conditions. The extraordinarily high rates of population
turnover and potentially highly selective feeding behavior of gastrotrichs suggest that they may exert a considerable influence on the composition of natural bacterial
communities.
D. Evolutionary Relationships
The phylogenetic relationships among the various
aschelminth groups are not yet clear. Several points
of morphological similarity between nematodes and
gastrotrichs (summarized by Lorenzen, 1985, and
Wallace et al., 1996) suggest that these two groups are
allied, but other analyses (Conway Morris, 1993;
Garey et al., 1996) have placed gastrotrichs closer to
rotifers and even platyhelminthes than to nematodes.
Within the Gastrotricha, only the broad outlines
of phylogeny have been sketched out. Three major
groups of gastrotrichs are widely recognized: the order
Macrodasyida and the suborders Multitubulata and
Paucitubulata of the order Chaetonotida. The Multitubulata (which includes only the marine Neodasys)
exhibit many primitive characteristics and are in some
ways intermediate between the Macrodasyida and
the Paucitubulata (which contains almost all of the
freshwater gastrotrichs). Therefore, Boaden (1985)
suggested that modern Macrodasyida and Paucitubulata are descended from a Neodasys-like ancestor.
Nevertheless, on the basis of digestive tract ciliature
and other characters, we believe that the ancestral
188
David Strayer and William D. Hummon
gastrotrich was more likely similar to the dactylopodolid macrodasyids.
Evolutionary relationships among the families,
genera, and species of freshwater gastrotrichs are still
largely unknown because of a paucity of basic morphological, biochemical, and zoogeographical information about most species. An enormous number of
species have not even been described, let alone studied.
Even in North America, perhaps 75 – 90% of the
species of freshwater gastrotrichs are undescribed. The
correct systematic placement of some genera (e.g.,
Dichaetura, Marinellina) will require much additional
study. Furthermore, there is growing concern (e.g.,
Remane, 1935 – 1936; Ruppert, 1977; Kisielewski,
1981, 1987, 1991, 1997; Schwank, 1990) that some
of the traditionally defined genera of the Chaetonotidae may be inadequate. Better morphological information and the discovery of species having characters
intermediate between genera is leading to a blurring of
distinctions between genera such as Chaetonotus,
Heterolepidoderma, and Lepidodermella. Further, it is
becoming increasingly clear that some of the characters traditionally used to define gastrotrich genera
(e.g., adhesive tubes, cuticle ornamentation) are subject to convergent evolution (e.g., Kisielewski, 1991;
Balsamo and Fregni, 1995), and so might not reflect
common lines of descent. Finally, it seems likely that
Ichthydium, which is defined by an absence of cuticular ornamentation, is polyphyletic. In response to these
concerns, Schwank (1990) and Kisielewski (1991)
erected new genera of chaetontotids and redefined
existing genera, but the classification of this group
may need to be redone once more complete information becomes available.
IV. COLLECTING, REARING, AND PREPARATION
FOR IDENTIFICATION
Gastrotrichs may be collected by taking samples of
sediments or vegetation. For quantitative work on sediment-dwelling species, small-diameter (2 – 5 cm) cores
are preferable. For plant-dwelling forms, quantitative
samples probably could be obtained by modifying sampling methods developed for macroinvertebrates (e.g.,
Downing, 1984; Jackson, 1997) to use very fine mesh
and small sample volumes or subsampling.
Because living animals are preferable to preserved
animals for many purposes, it often is desirable to extract the animals from the sample prior to preservation.
If the sample must be preserved immediately, workers
recommend narcotizing the animals with 1% MgCl2
for 10 min, then fixing them in 10% formalin with rose
bengal (e.g., Hummon, 1981, 1987).
It is difficult to extract or count the gastrotrichs
from a sample. Some workers have handpicked or
counted animals under a dissecting microscope (e.g.,
Hummon, 1981, 1987; Evans, 1982; Strayer, 1985),
but this procedure is tedious. Density gradient centrifugation (Nichols, 1979; Schwinghamer, 1981) ought to
be useful in extracting gastrotrichs from sediment, but
has not yet been tested on freshwater gastrotrichs.
Nesteruk (1987) found that a modified Baermann funnel was useful for extracting chaetonotids, but not
dasydytids, from pond sediments. Sieves should be
avoided or used cautiously, because gastrotrichs are too
small to be retained quantitatively on even very fine
mesh sieves. For example, Hummon (1981) found that
a 37-m mesh sieve retained only 31% of the gastrotrichs in a series of samples taken from the upper
Mississippi River. For quantitative work, it is important to check the efficiency of whatever extraction or
counting method is used, because gastrotrichs are so
small and can be easily overlooked (cf. Strayer, 1985,
pp. 295 – 296).
Living gastrotrichs are preferable to dead gastrotrichs for taxonomic work. Living gastrotrichs often
are too active for critical observations to be made, so
they must be slowed down. This can be accomplished
by gently squeezing the animal (either with a rotocompressor — Spoon, 1978 — or by removing some of the
water from beneath a cover slip with a tissue), by placing it in a viscous medium such as methylcellulose, or
by narcotizing it. Cocaine was the traditional narcotic
of choice (Brunson, 1959), but it is now difficult to
obtain for laboratory use. d’Hondt (1967) recommended using MS 222, and we have had very good
success with narcotizing gastrotrichs by bleeding 1.8%
neosynephrine (available at pharmacies) under the
cover slip. Animals may be killed with formalin or
fumes of osmium tetroxide (e.g., Brunson, 1950) following narcotization. Osmium tetroxide is a superior
fixative, but is dangerous and should be used with extreme care. It is sometimes necessary to examine individual scales, which can be isolated from an animal by
bleeding 2% acetic acid under the cover slip. For serious taxonomic work, videomicroscopy of living animals may provide better permanent documentation of
species characters than killed specimens or slides.
Procedures for culturing gastrotrichs were described by Packard (1936), Brunson (1949), Townes
(1968), Hummon (1974), and Bennett (1979). Gastrotrichs have been cultured on 0.1% malted milk, raw
egg yolk, wheat grain infusion, baked lettuce infusion,
and baker’s yeast. Brunson (1949) recommended that
animals be acclimated gradually to culture media when
collected from the wild. Hummon (1974) described a
procedure for starting individual cultures of known-age
7. Gastrotricha
animals from eggs that may be especially useful for
bioassay work.
V. TAXONOMIC KEY
It is relatively easy to identify most North American freshwater gastrotrichs to genus and very difficult
to identify them to species. Species identification requires a keen eye, careful observation, a cooperative
gastrotrich, and some luck, because most of the freshwater gastrotrichs of North America undoubtedly are
undescribed. The following works are helpful in species
identification: Brunson (1950, 1959), who keyed and
illustrated species then known from North American
189
freshwaters; Robbins (1965, 1973), who gave additional information and drawings of North American
species; d’Hondt (1971b), who gave a key to the
species of Lepidodermella and defined three subgenera
of Ichthydium; Kisielewski (1981), who made a critical
evaluation of the morphological characters that must
be measured to describe (or identify) a species;
Kisielewski (1986b), who gave a recent treatment of
Aspidiophorus; Schwank (1990), who provided illustrated keys (in German) for all known freshwater gastrotrichs worldwide; and Kisielewski (1991), who described many Brazilian species and addressed several
important issues in gastrotrich systematics. The following key includes genera of freshwater gastrotrichs
known from or likely to be found in North America.
1a.
Animal with at least three pairs of adhesive tubules (one anterior and two posterior) and a pair of pharyngeal pores, body strapshaped (Fig. 1A,B); an almost entirely marine group not yet reported from North American freshwaters.............order Macrodasyida
1b.
Animal lacking adhesive tubules (Fig. 7A– H ) or with one pair (very rarely two pairs) of adhesive tubules posteriorly (Fig. 6A– L),
and no pharyngeal pores, body strap-shaped or tenpin-shaped; common and widespread in freshwater ...............order Chaetonotida
...........................................................................................................................................................................................................2
2a (1).
Posterior end of body usually with furca and adhesive tubules; body usually strap-shaped or tenpin-shaped (Fig. 6A-L) ...................3
2b.
Posterior end of body without furca or adhesive tubules, although sometimes bearing spines or pegs; body usually tenpin or bottleshaped (Fig. 7A– H)..........................................................................................................................................................................12
3a (2).
Furca doubly branched; scales and spines sparse or absent (Fig. 6A); a rare genus not yet reported from North America ....................
Dichaeturidae, Dichaetura
3b.
Furca singly branched; scales and spines present or absent (Fig. 6B– L); common and widespread .....................................................4
4a (3).
Branches of furca heavy, sickle-shaped, curved, and tapered, not distinctly divided into a cone-shaped basal part and a distal duct;
head with long cilia that are not arranged in bundles; head plates absent (Fig. 6B); a rare family not yet reported from North America ................................................................................................................................................................................Proichthydiidae
4b.
Branches of furca usually with a cone-shaped base and a distal adhesive duct; body often with numerous spines or scales; head with
cilia that are arranged in bundles; cephalic plates present (Fig. 6C – L); common and widespread ................................Chaetonotidae
...........................................................................................................................................................................................................5
5a (4).
Branches of furca ringed and often very long; body often large and without a distinct neck (Fig. 6C) ..............................Polymerurus
5b.
Branches of furca not ringed (Fig. 6D – L)...........................................................................................................................................6
6a (5).
Dorsal surface of body without spines or scales (except for dorsal sensory bristles or a few scales at the base of the furca (Fig. 6D) ...
...........................................................................................................................................................................................Ichthydium
6b.
Dorsal surface of body with numerous scales or spines (Fig. 6E – L) ...................................................................................................7
7a (6).
Spines or spined scales present and often numerous (Fig. 6E – G, J) ....................................................................................................8
7b.
Spines absent (occasionally a few thin spines are present at the base of the furca) (Fig. 6H,I ,K,L) .....................................................9
8a. (7).
Ventral scales different from dorsal scales; spines of various types (Fig. 6E – G); common and widespread.......................Chaetonotus
8b.
Ventral scales similar to dorsal scales; posterior part of body with several long spines that reach beyond the end of the furca
(Fig. 6J); not yet reported from North America .............................................................................................................Lepidochaetus
9a (7).
Furca without adhesive tubes; body markedly tenpin-shaped, with groups of long cilia on the head and posterior part of the body
(Fig. 6H); a semiplanktonic genus known only from Brazil ......................................................................................................Undula
9b.
Furca with adhesive tubes; body strap-shaped or tenpin-shaped; without groups of long cilia on head and posterior body
(Fig. 6I,K,L) .....................................................................................................................................................................................10
10a (9).
Dorsal surface of body covered with stalked scales (Fig. 6I) ..........................................................................................Aspidiophorus
190
David Strayer and William D. Hummon
FIGURE 6
Genera of freshwater Dichaeturidae, Proichthydiidae, and Chaetonotidae. (A) Dichaetura;
(B) Proichthydium; (C) Polymerurus, showing detail of ringed branches of furca; (D) Ichthydium, (E – G)
Chaetonotus, showing examples of spination; (H) Undula; (I) Aspidiophorus, with detail of coat of scales and
a single scale; (J) Lepidochaetus, in dorsal (left) and ventral (right) views, (K) Heterolepidoderma, with detail
of scales; and (L) Lepidodermella, with detail of scales. From Remane (1935 – 1936), Brunson (1950), Hyman
(1951), Voigt (1958), Robbins (1965), and Kisielewski (1991).
10b.
Scales on dorsal surface of body unstalked (Fig. 6K,L) .....................................................................................................................11
11a (10).
Scales elongate, with longitudinal keels (Fig. 6K) ...................................................................................................Heterolepidoderma
11b.
Scales not keeled (Fig. 6L)............................................................................................................................................Lepidodermella
12a (2).
Head with club-shaped tentacles (Fig. 7A,B) .............................................Neogosseidae..................................................................13
12b.
Head without club-shaped tentacles (Fig. 7C – H) .......................................Dasydytidae..................................................................14
13a (12).
Posterior end of body with two groups of long spines (Fig. 7A); elements of mouth-ring jointed.........................................Neogossea
13b.
Posterior end of body with single medial group of spines (Fig. 7B); elements of mouth-ring unjointed .............................Kijanebalola
7. Gastrotricha
191
FIGURE 7
Genera of freshwater Neogosseidae and Dasydytidae. (A) Neogossea; (B) Kijanebalola; (C)
Stylochaeta; (D) Ornamentula; (E) Anacanthoderma; (F) Setopus; (G) Haltidytes, and (H) Dasydytes. From
Brunson (1950), Krivanek and Krivanek (1958), Balsamo (1983), Schwank (1990), and Kisielewski (1991).
14a (12).
Posterior end of body with pair of peglike protuberances (Fig. 7C) ....................................................................................Stylochaeta
14b.
Posterior end of body without peglike protuberances (Fig. 7D – H) ..................................................................................................15
15a (14).
Body enclosed in a “lorica” of large, thick, ornamented scales (Fig. 7D); known only from Brazil..................................Ornamentula
15b.
Scales absent or small and inconspicuous (Fig. 7E – H) .....................................................................................................................16
16a (15).
Head much narrower than body and scarcely wider than neck; lateral spines absent or identical to dorsal spines; pharynx with two
bulbs (Fig. 7E); not yet reported from North America ...............................................................................................Anacanthoderma
16b.
Head distinctly wider than neck; body with long lateral spines; pharynx with one bulb or no bulb (Fig. 7F – H) .............................17
17a (16).
Some lateral spines movable and sharply bent basally; posterior end of body rounded, without rear spines (Fig. 7G)..........Haltidytes
17b.
All spines fixed, straight or bent distally; posterior end of body usually with spines (Fig. 7F,H).......................................................18
18a (17).
Lateral spines with 1 – 3 lateral denticles and often terminally bifurcated; spines of uniform thickness from their base to the last lateral denticle; pharynx usually with a distinct posterior bulb (Fig. 7H) .................................................................................Dasydytes
18b.
Lateral spines tapered, with at most one weak lateral denticle and never terminally bifurcated; pharynx without posterior bulb
(Fig. 7F) ...................................................................................................................................................................................Setopus
192
David Strayer and William D. Hummon
VI. LITERATURE CITED
Balsamo, M. 1980. Spectral sensitivity in a freshwater gastrotrich
(Lepidodermella squammatum Dujardin). Experientia 36:
830 – 831.
Balsamo, M. 1981. Gastrotrichi della Toscana: il lago di Sibolla.
Bolletino del Museo Civico di Storia Naturale Verona 7:
547 – 571.
Balsamo, M. 1983. Gastrotrichi (Gastrotricha). Guide per il Riconoscimento delle Specie Animali delle Acque Interne Italiane
20. Consiglio Nazionale delle Ricerche.
Balsamo, M., Fregni, E. 1995. Gastrotrichs from interstitial freshwater, with a description of four new species. Hydrobiologia
302:163 – 175.
Bennett, L. W. 1979. Experimental analysis of the trophic ecology of
Lepidodermella squammata (Gastrotricha:Chaetonotida) in
mixed culture. Transactions of the American Microscopical
Society 98:254 – 260.
Bertolani, R., Balsamo, M. 1989. Tardigradi e Gastrotrichi del
Trentino: il Lago di Tovel. Studi Trentini di Scienze Naturali
65:83 – 93.
Blinn, D. W., Green, J. 1986. A pump sampler study of microdistribution in Walker Lake, Arizona, USA.: a senescent crater lake.
Freshwater Biology 16:175 – 185.
Boaden, P. J. S. 1985. Why is a gastrotrich?, in: Conway Morris, S.,
George, J. D., Gibson, R., Platt, H. M. Eds., The origins and relationships of lower invertebrates. Clarendon Press, Oxford,
UK, pp. 248 – 260.
Bovee, E. C., Cordell, D. L. 1971. Feeding on gastrotrichs by the
heliozoon Actinophrys sol. Transactions of the American
Microscopical Society 90:365 – 369.
Brandenburg, J. 1962. Elektronenmikroscopische Untersuchungen
des Terminalapparates von Chaetonotus sp. (Gastrotrichen) als
Beispeil einer Cyrtocyte bei Aschelminthen. Zeitschrift für
Zellforschung 57:136 – 144.
Brunson, R. B. 1947. Gastrotricha of North America. II. Four new
species of Ichthydium from Michigan. Transactions of the
Michigan Academy of Science, Arts, and Letters 33:59 – 62.
Brunson, R. B. 1948. Chaetonotus tachyneusticus: a new species
of gastrotrich from Michigan. Transactions of the American
Microscopical Society 67:350 – 351.
Brunson, R. B. 1949. The life history and ecology of two North
American gastrotrichs. Transactions of the American Microscopical Society 68:1 – 20.
Brunson, R. B. 1950. An introduction to the taxonomy of the Gastrotricha with a study of eighteen species from Michigan. Transactions of the American Microscopical Society 69:325 – 352.
Brunson, R. B. 1959. Gastrotricha, in: Edmondson, W. T. Ed., Freshwater biology. 2nd ed., Wiley, New York, pp. 406 – 419.
Cole, G. A. 1955. An ecological study of the microbenthic fauna of two
Minnesota lakes. American Midland Naturalist 53:213 – 230.
Conway Morris, S. 1993. The fossil record and the early evolution of
the Metazoa. Nature 361:219 – 225.
Craft, L. 1993. A taxonomic survey of freshwater gastrotrichs from
eastern Ohio. MS thesis, Ohio University, 169 pp.
Davis, K. B. 1937. The Gastrotricha (Chaetonotoidea) of Washington
County, Arkansas. MA thesis, University of Arkansas, 94 pp.
Downing, J. A. 1984. Sampling the benthos of standing waters, in:
Downing, J. A., Rigler, F. H. Eds., A manual on methods for the
assessment of secondary productivity in freshwaters. 2nd ed.,
Blackwell Scientific Publications, Oxford, UK, pp. 87–130
Emberton, K. C. 1981. First record of Chaetonotus heideri
(Gastrotricha: Chaetonotidae) in North America. Ohio Journal
of Science 81:95 – 96.
Evans, W. A. 1982. Abundances of micrometazoans in three sandy
beaches in the island area of western Lake Erie. Ohio Journal of
Science 82:246 – 251.
Faucon, A. S., Hummon, W. D. 1976. Effects of mine acid on the
longevity and reproductive rate of the Gastrotricha Lepidodermella squammata (Dujardin). Hydrobiologia 50:265 – 269.
Frey, D. G. 1982. Questions concerning cosmopolitanism in Cladocera. Archiv für Hydrobiologie 93:484 – 502.
Garey, J. R., Krotec, M., Nelson, D. R., Brooks, J. 1996. Molecular
analysis supports a tardigrade-arthropod association. Invertebrate Biology 115:79 – 88.
Gray, J. S., Johnson, R. M. 1970. The bacteria of a sandy beach as
an ecological factor affecting the interstitial gastrotrich
Turbanella hyalina Schultze. Journal of Experimental Marine
Biology and Ecology 4:119 – 133.
Green, J. 1986. Associations of zooplankton in six crater lakes in
Arizona, Mexico and New Mexico. Journal of Zoology
(A)208:135 – 159.
d’Hondt, J.-L. 1967. Effets de quelques anesthétiques sur les Gastrotriches. Experientia 23:1025 – 1026.
d’Hondt, J.-L. 1971a. Gastrotricha. Oceanography and Marine
Biology: Annual Review 9:141 – 192.
d’Hondt, J.-L. 1971b. Note sur quelques Gastrotriches Chaetonotidae. Bulletin de la Société Zoologique de France 96:215 – 235.
Horlick, R. G. 1975. Dasydytes monile, a new species of gastrotrich
from Illinois. Transactions of the Illinois State Academy of
Science 68:61 – 64.
Hummon, M. R. 1984a. Reproduction and sexual development in a
freshwater gastrotrich. 1. Oogenesis of parthenogenic eggs
(Gastrotricha). Zoomorphology 104:33 – 41.
Hummon, M. R. 1984b. Reproduction and sexual development in a
freshwater gastrotrich. 2. Kinetics and fine structure of postparthenogenic sperm formation. Cell and Tissue Research
236:619 – 628.
Hummon, M. R. 1984c. Reproduction and sexual development in a
freshwater gastrotrich. 3. Postparthenogenic development of
primary oocytes and the X-body. Cell and Tissue Research
236:629 – 636.
Hummon, M. R. 1986. Reproduction and sexual development in a
freshwater gastrotrich. 4. Life history traits and the possibility
of sexual reproduction. Transactions of the American Microscopical Society 105:97 – 109.
Hummon, M. R., Hummon, W. D. 1979. Reduction in fitness of the
gastrotrich Lepidodermella squammata by dilute mine acid
water and amelioration of the effect by carbonates. International Journal of Invertebrate Reproduction 1:297 – 306.
Hummon, M. R., Hummon, W. D. 1983. Gastrotricha, in: Adiyodi
K. G., Adiyodi, R. G. Eds., Reproductive biology of invertebrates. Vol II: Spermatogenesis and sperm function. Wiley, London, pp. 195 – 205.
Hummon, W. D. 1974. Effects of DDT on longevity and reproductive
rate in Lepidodermella squammata (Gastrotricha, Chaetonotida).
American Midland Naturalist 92:327 – 339.
Hummon, W. D. 1981. Extraction by sieving: a biased procedure in
studies of stream meiobenthos. Transactions of the American
Microscopical Society 100:278 – 284.
Hummon, W. D. 1982. Gastrotricha, in: Parker, S. P. Ed., Synopsis
and classification of living organisms. Vol. 1. McGraw – Hill,
New York, pp. 857 – 863.
Hummon, W. D. 1987. Meiobenthos of the Mississippi headwaters, in: Bertolani, R. Ed., Biology of tardigrades. Selected
Symposia and Monographs UZI., 1, Mucchi, Modena, Italy,
pp. 125 – 140.
Hummon, W. D., Hummon, M. R. 1983. Gastrotricha, in: Adiyodi,
K. G., Adiyodi, R. G. Eds., Reproductive biology of inverte-
7. Gastrotricha
brates. Vol. I: Oogenesis, oviposition, and oosorption. Wiley,
London, pp. 211 – 221.
Hummon, W. D., Hummon, M. R. 1988. Gastrotricha, in: Adiyodi,
K. G., Adiyodi, R. G. Eds., Reproductive biology of invertebrates. Vol. III: Accessory sex glands. Oxford and IBH Publishing Company, New Delhi, pp. 81 – 85.
Hummon, W. D., Evans, W. A., Hummon, M. R., Doherty, F. G.,
Wainberg, R. H., Stanley, W. S. 1978. Meiofaunal abundance in
sandbars of acid mine polluted, reclaimed, and unpolluted
streams in southeastern Ohio, in: Thorp, J. H., Gibbons, J. W.
Eds., Energy and environmental stress in aquatic ecosystems.
DOE Symposium Series (CONF – 771114). National Technical
Information Service, Springfield, VA, pp.188 – 203.
Hutchinson, G. E. 1967. A treatise on limnology, Vol 2: Introduction
to lake biology and the limnoplankton. Wiley, New York.
Hyman, L. H. 1951. The invertebrates: Acanthocephala, Aschelminthes, and Entoprocta: the pseudocoelomate Bilateria.
Vol. 3. McGraw-Hill, New York.
Jackson, M. J. 1997. Sampling methods for studying macroinvertebrates in the littoral vegetation of shallow lakes. The Broads
Authority, Norwich, UK.
Kisielewska, G. 1981. Hermaphroditism of freshwater gastrotrichs in
natural conditions. Bulletin de l’Académie Polonaise des Sciences, Série des Sciences Biologiques 29:167 – 172.
Kisielewska, G. 1982. Gastrotricha of two complexes of peat hags
near Siedlce. Fragmenta Faunistica 27:39 – 57.
Kisielewski, J. 1981. Gastrotricha from raised and transitional peat
bogs in Poland. Monografie Fauny Polski 11:1 – 143.
Kisielewski, J. 1986a. Freshwater Gastrotricha of Poland. VII. Gastrotricha of extremely eutrophicated water bodies. Fragmenta
Faunistica 30:267 – 295.
Kisielewski, J. 1986b. Taxonomic notes on freshwater gastrotrichs of
the genus Aspidiophorus Voigt (Gastrotricha: Chaetonotidae),
with descriptions of four new species. Fragmenta Faunistica
30:139 – 156.
Kisielewski, J. 1987. Two new interesting genera of Gastrotricha
(Macrodasyida and Chaetonotida) from the Brazilian freshwater psammon. Hydrobiologia 153:23 – 30.
Kisielewski, J. 1991. Inland-water Gastrotricha from Brazil. Annales
Zoologici 43 (Suppl 2):1 – 168.
Kisielewski, J. 1997. On the subgeneric division of the genus Chaetonotus Ehrenberg (Gastrotricha). Annales Zoologici 46:145 – 151.
Kisielewski, J. 1998. Brzuchorzeski (Gastrotricha). Fauna Slodkowodna Polski 31. Polskie Towarzystwo Hydrobiologiczne,
Uniwersytet Lodzki.
Krivanek, R. C., Krivanek, J. O. 1958. A new and a redescribed
species of Neogossea (Gastrotricha) from Louisiana. Transactions of the American Microscopical Society 77:423 – 428.
Levy, D. P. 1984. Obligate postparthenogenetic hermaphroditism and
other evidence for sexuality in the life cycles of freshwater Gastrotricha. Dissertation, Rutgers University, New Brunswick, NJ.
257 pp.
Levy, D. P., Weiss, M. J. 1980. Sperm in the life cycle of the freshwater gastrotrich, Lepidodermella squammata. American Zoologist 20:749 (abstract).
Lorenzen, S. 1985. Phylogenetic aspects of pseudocoelomate evolution, in: Conway Morris, S., George, J. D., Gibson, R., Platt,
H.M. Eds., The origins and relationships of lower invertebrates.
Clarendon Press, Oxford, pp. 210 – 223.
Madoni, P. 1989. Community structure of the microzoobenthos in
Lake Suviana (Tusco-Emilian Apennines). Bolletino di Zoologia
56:159 – 165.
Moore, G. M. 1939. A limnological investigation of the microscopic
benthic fauna of Douglas Lake, Michigan. Ecological Monographs 9:537 – 582.
193
Moore, J. W. 1979. Some factors influencing the distribution, seasonal abundance and feeding of subarctic Chironomidae
(Diptera). Archiv für Hydrobiologie 85:302 – 325.
Nesteruk, T. 1986. Freshwater Gastrotricha of Poland. IV. Gastrotricha from fish ponds in the vicinity of Siedlce. Fragmenta
Faunistica 30:215 – 233.
Nesteruk, T. 1987. Assessing the efficiency of three methods of extracting freshwater Gastrotricha from bottom mud. Acta Hydrobiologica 29:219 – 226.
Nesteruk, T. 1991. Vertical distribution of Gastrotricha in organic
bottom sediment of inland water bodies. Acta Hydrobiologica
33:253 – 264.
Nesteruk, T. 1996. Density and biomass of Gastrotricha in sediments
of different types of standing waters. Hydrobiologia 324:
205 – 208.
Nichols, J. A. 1979. A simple flotation technique for separating
meiobenthic nematodes from fine-grained sediments. Transactions of the American Microscopical Society 98:127 – 130.
Nuss, B. 1984. Ultrastrukturelle und ökophysiologische Untersuchungen an kristalloiden Einschlüssen der Muskeln eines
sulfidtoleranten limnischen Nematoden (Tobrilus gracilis).
Veröffentlichungen des Instituts für Meeresforschung in Bremerhaven 20:3 – 15.
Nuss, B., Trimkowski, V. 1984. Physikalische Mikroanalysen an
kristalloiden Einschlüssen bei Tobrilus gracilis (Nematoda, Enoplida). Veröffentlichungen des Instituts für Meeresforschung in
Bremerhaven 20:17 – 27.
Packard, C. E. 1936. Observations on the Gastrotricha indigenous to
New Hampshire. Transactions of the American Microscopical
Society 55:422 – 427.
Pennak, R. W. 1978. Freshwater invertebrates of the United States.
2nd ed., Wiley-Interscience, New York.
Pfaltzgraff, G. H. 1967. A preliminary study of the Gastrotricha of
northern Indiana. Proceedings of the Indiana Academy of
Sciences 76:400 – 404.
Powell, E. N., Crenshaw, M. A., Rieger, R. M. 1979. Adaptations to
sulfide in the meiofauna of the sulfide system. I. 35Sulfide accumulation and the presence of a sulfide detoxification system.
Journal of Experimental Marine Biology and Ecology
37:57 – 76.
Powell, E. N., Crenshaw, M. A., Rieger, R. M. 1980. Adaptations to
sulfide in the sulfide-system meiofauna. Endproducts of sulfide
detoxification in three turbellarians and a gastrotrich. Marine
Ecology Progress Series 2:169 – 177.
Preobrajenskaja, E. N. 1926. Zur Verbreitung der Gastrotrichen
in den Gewässern der Umgebung zu Kossino (in Russian, with
a German summary). Arbeiten der biologischen Station zu
Kossino (bei Moskau) 4:3 – 14.
Remane, A. 1935 – 1936. Gastrotricha und Kinorhyncha. Klassen
und Ordnungen des Tierreichs 4 (Abteilung 2, Buch 1, Teil 2,
Lieferung 1 – 2):1 – 242, 373 – 385.
Remane, A. 1961. Neodasys uchidai nov. spec., eine zweite NeodasysArt (Gastrotricha Chaetonotida). Kieler Meeresforschungen
17:85 – 88.
Renaud-Mornant, J. 1986. Gastrotricha, in: Botosaneanu, L. Ed.,
Stygofauna mundi. E. J. Brill, Leiden, The Netherlands, pp.
86 – 109.
Robbins, C. E. 1965. Two new species of Gastrotricha (Aschelminthes) from Illinois. Transactions of the American Microscopical Society 84:260 – 263.
Robbins, C. E. 1973. Gastrotricha from Illinois. Transactions of the
Illinois State Academy of Science 66:124 – 126.
Rudescu, L. 1968. Die Rotatorien, Gastrotrichen und Tardigraden der
Schilfrohrgebiete des Donaudeltas. Hidrobiologia (Bucharest)
9:195 – 202.
194
David Strayer and William D. Hummon
Ruppert, E. E. 1977. Ichthydium hummoni n. sp., a new marine
chaetonotid gastrotrich with a male reproductive system.
Cahiers de Biologie Marine 18:1 – 5.
Ruppert, E. E. 1988. Gastrotricha, in: Higgins, R. P., Thiel, H. Eds.,
Introduction to the study of meiofauna. Smithsonian Institution
Press, Washington, pp. 302 – 311.
Ruttner-Kolisko, A. 1955. Rheomorpha neiswestnovae und Marinellina
flagellata, zwei phylogenetisch interessante Wurmtypen aus
dem Süsswasserpsammon. Österreichische Zoologische Zeitschrift
6:55 – 69.
Schmid-Araya, J. M., Schmid, P. E. 1995. The invertebrate species of
a gravel stream. Jahresberichte der Biologischen Station Lunz
15:11 – 21.
Schwank, P. 1990. Gastrotricha, in: Schwoerbel, J., Zwick, P. Eds.,
Süsswasserfauna von Mitteleuropa, Band 3. Gustav Fischer Verlag, Stuttgart, pp. 1 – 252.
Schwinghamer, P. 1981. Extraction of living meiofauna from marine
sediments by centrifugation in a silica sol – sorbitol mixture.
Canadian Journal of Fisheries and Aquatic Sciences 38:476 – 478.
Spoon, D. M. 1978. A new rotary microcompressor. Transactions of
the American Microscopical Society 97:412 – 416.
Strayer, D. 1985. The benthic micrometazoans of Mirror Lake, New
Hampshire. Archiv für Hydrobiologie Supplementband 72:
287 – 426.
Sudzuki, M. 1971. Die das Kapillarwasser des Lückensystems bewohnenden Gastrotrichen Japans I. Zoological Magazine 80:
256 – 257.
Todaro, M. A., Fleeger, J. W., Hu, Y. P., Hrincevich, A.W., Foltz, D.
W. 1996. Are meiofaunal species cosmopolitan? Morphological
and molecular analysis of Xenotrichula intermedia (Gastrotricha: Chaetonotida). Marine Biology 125:735 – 742.
Townes, M. M. 1968. The collection, identification, and cultivation
of gastrotrichs. Turtox News 46:99 – 101.
Voigt, M. 1958. Gastrotricha. Die Tierwelt Mitteleuropas, Band I,
Lieferung 4a, pp. 1 – 45.
Wallace, R. L., Ricci, C., Melone, G., 1996. A cladistic analysis of
pseudocoelomate (aschelminth) morphology. Invertebrate Biology 115:104 – 112.
Weiss, M. J., Levy, D. P. 1979. Sperm in “parthenogenetic” freshwater gastrotrichs. Science 205:302 – 303.
8
PHYLUM ROTIFERA
Robert Lee Wallace
Terry W. Snell
Department of Biology
Ripon College
Ripon, Wisconsin 54971
School of Biology
Georgia Institute of Technology
Atlanta, Georgia 30332
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Organ-System Function
C. Environmental Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Collecting, Rearing, and Preparation
for Identification
I. INTRODUCTION
The phylum Rotifera or Rotatoria comprises of approximately 2000 species of unsegmented, bilaterally
symmetrical, pseudocoelomates, possessing two distinctive features (Fig. 1). First, at the apical end (head) is a
ciliated region called the corona, which is used in locomotion and food gathering. In adults of some forms,
ciliation is lacking and the corona is a funnel or bowlshaped structure at the bottom of which is the mouth.
Second, a muscular pharynx, the mastax, possessing a
complex set of hard jaws, called trophi, is present in all
rotifers.
When viewing the anterior end of most rotifers one
is struck with the idea of a rotating wheel. This is due
to the metachronal beat of cilia on the corona, a structure usually composed of two concentric rings: trochus
and cingulum (Fig. 2). This same image provided early
microscopists with the name for the phylum: the etymon is Latin, rota, “wheel” and Latin, ferre, “to bear”
equals “wheel bearers.” Although rotifers are often
confused with ciliated protozoans and gastrotrichs by
beginning students, those organisms do not possess
trophi and their ciliation is not distributed in the same
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
A. Collections
B. Culture
C. Preparation for Identification
V. Classification and Systematics
A. Classification
B. Systematics
C. Taxonomic Keys
D. Taxonomic Key to Families
of Freshwater Rotifera
Literature Cited
way as in rotifers. Rotifers are small organisms, generally ranging from 100 – 1,000 m long, although a few
elongate species may surpass 2,000 m or more. Very
few rotifers are parasitic (May, 1989); nearly all are
free-living herbivores or predators.
Collectively this phylum is widely distributed, being found in all freshwater habitats at densities generally ranging up to about 1,000 individuals/L. However,
rotifers occasionally become abundant if sufficient food
is available, and can attain population densities of
5,000 individuals/L. In some rather unusual water
bodies, exceedingly large populations can develop;
sewage ponds may contain about 12,000 individuals/L
(Seaman et al., 1986), and at certain times in soda water bodies in Chad, much more than 100,000 individuals/L may occur (Iltis and Riou-Duvat, 1971)! Although most inhabit freshwaters, some genera also
have members that occur in brackish and marine waters. For example, about 20 of the 32 species comprising the genus Synchaeta are described as marine (Nogrady, 1982). However, only about 50 species of
rotifers are exclusively marine. In general, rotifers are
not as diverse or as abundant in marine environments
as microcrustaceans, but they occur in many nearshore
195
196
Robert Lee Wallace and Terry W. Snell
FIGURE 1 Lateral view of a generalized rotifer. (Modified from
Koste and Shiel, 1987, with permission.)
marine communities (Egloff, 1988) and occasionally
comprise the dominant portion of the biomass
(Schnese, 1973; Johansson, 1983). One unusual group
of rotifers, the bdelloids (Fig. 3), may be found inhabiting the film of water covering mosses, lichens, and liverworts. Additionally, they are often abundant in soils
(Pourriot, 1979); estimates of their densities range from
about 32,000 to more than 2 million individuals/m2,
depending on soil moisture levels. Because of their
feeding habits, and the fact that they are sometimes
more numerous than nematodes, rotifers play an important role in nutrient cycling in soils (Pourriot,
1979).
Most rotifers are free-moving, either swimming as
members of the plankton or crawling over plants or
within the sediments; however, some sessile species live
permanently attached to freshwater plants (Wallace,
1980). The vast majority of rotifers are solitary, but
about 25 species form colonies of various sizes (Wallace, 1987). All freshwater rotifers are either exclusively parthenogenetic or produce males for a limited
time each year. Therefore, unless collections are made
FIGURE 2 Female and male Brachionus plicalitis. (Modified from Pourriot, 1986, with permission.)
8. Phylum Rotifera
197
ologia. Some of the papers discussed in this chapter
were presented at those meetings.
II. ANATOMY AND PHYSIOLOGY
A. External Morphology
FIGURE 3 Typical bdelloid rotifer (Philodina). (Modified from
several sources.)
frequently, male rotifers may never be seen. Three very
different classes of rotifers are commonly recognized
(Seisonidea, Bdelloidea, Monogononta).
Additional accounts of this phylum may be found
in most texts of general and invertebrate zoology, and
in some specialized books about freshwaters (Edmondson, 1959, pp. 420 – 494; Hutchinson, 1967, pp.
506 – 551; Pennak, 1989, pp. 169 – 225). For detailed
reviews of the biology of rotifers consult the works of
de Beauchamp (1965), Hyman (1951, pp. 59 – 151),
Koste (1978), Ruttner-Kolisko (1974), and Nogrady
et al. (1993). In the 1800s there were some beautifully
illustrated works that still offer an excellent view of
these animals (e.g., Hudson and Gosse, 1886). There
is no single scientific journal or set of journals in
which researchers publish their work on rotifers;
the field is simply too diverse. However, every three
years, since 1976, a small group of workers (approximately 50 – 100) have gathered to hold the International Rotifer Symposium. To date, nine such meetings
have been held and most of the proceedings have been
published as a special volume of the journal Hydrobi-
Rotifers are saccate-to-cylindrical in shape, sometimes appearing wormlike (e.g., many bdelloids, Fig. 3).
Typically, the body is comprised of four regions: head
(with corona); neck; body; and foot (Fig. 1). Although
these regions may be marked by folds in the body wall,
which function like joints, rotifers are not metameric
(segmented). Unfortunately, the generalizations noted
here do not represent all rotifers well. In some forms,
the neck and foot may be quite prominent, while in others, they are absent (Fig. 4). In the class Monogononta,
male rotifers (Fig. 2) and juveniles (larval females) of
sessile rotifers are usually much smaller than adult females (Fig. 5). In addition, male rotifers are often structurally simpler (e.g., the gut commonly functions only
as a food reserve) and the larvae of sessile forms generally have a different morphology from that of the adult
(Wallace, 1980). When either males or larvae are found
in plankton tows, their strikingly different morphologies may lead to improper identification.
The foot is an appendage that extends from the
body ventrally (Fig. 2). It usually possesses two toes,
but the number may vary from 0 to 4. The foot also
may possess pedal glands whose ducts exit near the
toes. These glands secrete a sticky cement for temporarily attaching the rotifer to substrates. In larvae of
sessile rotifers, the cement forms a bond with the
substrate that is not easily detached; if dislodged, sessile forms do not reattach (Wallace, 1980).
Rotifers possess a syncytial integument or body
wall containing a filament layer of varying thickness
called the intracytoplasmic lamina (Clément, 1985).
This feature apparently is shared with the parasitic
phylum, Acanthocephala, indicating a phylogenetic
relationship (see Section III.D.). The integument of a
halophile rotifer, Brachionus plicatilis, was examined
biochemically by Bender and Kleinow (1988) and their
work indicates that it contains two filamentous, keratinlike proteins (Mr 39,000 and 47,000) cross-linked
by disulfide bonds. Species in which major portions of
the integument are thickened are termed “loricate;”
those in which the integument is thin and very flexible
are termed “illoricate.” In general, thickness of the
body wall is of little taxonomic significance since extremes may be found within a single family or genus
(e.g., Cephalodella). Further, even in loricate forms
portions of the integument (lorica) have a less welldeveloped intracytoplasmic lamina, making the lorica
198
Robert Lee Wallace and Terry W. Snell
FIGURE 4 Representative rotifers. Monogononts: (A) Asplanchna; (B) Euchlanis; (C) Keratella (lateral view);
(D) Polyarthra; and (E) Synchaeta. Bdelloids: (F) Philodina; and (G) Rotaria. (From Koste, 1976, with
permission.)
flexible in that region. Flexibility is found in the region
of the corona, often in the foot, and at articulations between movable spines and the body. In some rotifers,
the integument has various projections (e.g., bumps
and fixed or movable spines) that serve various functions, chief among these being protection from some
predators (see Sections III.A.1 and III.C.4).
B. Organ-System Function
Internally, rotifers possess a perivisceral cavity
(pseudocoelom) in which are suspended muscles, nerves,
and digestive, reproductive, and protonephridial organs
(Figs. 1, 2, and 3). Respiratory and circulatory systems
are absent. Rotifers possess two remarkable features.
First, all postembryonic tissues are syncytial. Second, all
individuals of a species have a consistent number of
nuclei in each organ throughout life. [N.B.: There are
approximately 900 nuclei per female (Hyman, 1951).]
This feature, called “eutely,” is also seen in a few other
invertebrates (e.g., nematodes, Chapter 9). Most nuclei
may be seen using a standard light microscope, but special optics such as differential interference contrast
(Nomarski) enhance their visibility.
1. Corona
There is considerable variation in the shape of the
anterior ends of rotifers, and at least seven different
8. Phylum Rotifera
199
FIGURE 5
Adults and larvae of some sessile species (family Flosculariidae). (A) Adult Octotrocha speciosa
(from Koste, 1989, with permission); (B) larval Octrotrocha speciosa (dark field illumination, length 1200
m); (C) colony of adult Lacinularia flosculosa (bright field illumination, individual length 1200 m); and
(D) larval Lacinularia flosculosa (bright field illumination, larvae somewhat compressed by the cover glass,
length 450 m).
types of coronae have been described based on the
placement of the mouth and the distribution of cilia
(Koste and Sheil, 1987). The corona form of many rotifers is comprised of two ciliated rings called the
trochus and cingulum (Figs. 2 and 5). These structures
are responsible for the production of water currents
that are used in locomotion and feeding. However, not
all rotifers possess this coronal ciliation. Adults of the
family Collothecidae exhibit the most extreme varia-
tion from the typical plan; in most collothecids, cilia
are nearly or completely lacking from the corona and
long setae surround the rim of a funnel-shaped structure known as the infundibulum (Latin, a funnel)
(Fig. 6). These setae prevent escape of prey when the
edges of the infundibulum fold over the victim, capturing it in a fashion similar to the Venus flytrap. However, some adult atrochidae possess no setae on their
corona (e.g. Cupelopagis). In other groups, ciliation
200
Robert Lee Wallace and Terry W. Snell
FIGURE 6 Three representatives of the order Collothecacea. (A) Collotheca trilobata, a sessile form; (B) Collotheca mutabilis, a planktonic form; (C) Collotheca sp. with two developing embryos (width of corona 100
m); and (D) Collotheca cornuta, dark field illumination (length of setae 200 m). (A, modified from
Wallace et al., 1989, with permission; and B, modified after several sources.)
8. Phylum Rotifera
may be limited to a ventral field or several lobes, as is
seen in some creeping forms and some bdelloids. Other
structures that may be present on the corona include
cirri, sensory antennae, and palps.
2. Trophi and Gut
Once food is captured by the corona, it enters a
ventral mouth by passing through a short, ciliated tube
into a muscular pharynx, termed the “mastax.” The
mastax possesses a chitinous lining on the inside, developed as a set of translucent jaws called trophi. The
trophi may work the food in various ways (e.g., grinding) before it is passed to the esophagus and swallowed
(Figs. 1 and 2). In the family Collothecidae, a portion
of the mastax is enlarged into a food-storage organ
known as the proventriculus (Fig. 6). The mastax leads
posteriorly to an esophagus, stomach, and in most
species, intestine and anus, but the gut ends in a blind
stomach in some genera (i.e., Asplanchna, Asplanchnopus). The posterior portion of the intestine (cloaca) receives eggs from the oviduct and fluid from either a
bladder or directly from paired protonephridia (Fig. 1).
Often the gut is pigmented, depending on the nature of
recently ingested material. Different species found in
the same sample may possess guts that vary in color
due to differences in diet.
Rotifer trophi are composed of several hard parts
and associated musculature, which articulate in a specific spatial arrangement. In their basic form, trophi
consist of three functional units: an incus (Latin, anvil)
and paired mallei (Latin, hammer) (Fig. 7). The incus is
composed of three pieces: a fulcrum and a pair of rami
(Latin, branch) that move like forceps and articulate
with the fulcrum at their bases. Each malleus consists of
two parts: manubrium and uncus. The manubrium
(Latin, handle) resembles a club-shaped structure extended at one end into a cauda (Latin, tail) and flared at
the other end (head). The manubrium articulates with a
toothed structure called the uncus (Latin, hook). The
plane of movement of the pieces comprising the malleus
is at right angles to that of the rami. In some species,
the trophi may be modified by reduction of the basic
parts, addition of accessory structures, or by asymmetrical development of one or more of the pieces.
The trophi of rotifers have been recognized as important taxonomic features: classes, orders, and families, and even species may be determined based on the
details of the trophi alone (e.g., Salt et al., 1978). Nine
different types of trophi are recognized based on the
size and shape of the seven pieces and the presence of
any accessory parts (Figs. 7 – 10), transitional and aberrant types also are known. In “Malleate” trophi (Fig.
8A – C, E, F, P), all parts of the incus and mallei are
well developed and functional, but the rami are charac-
201
teristically massive and may possess teeth along the inner margin. Further, the unci have 4 – 7 large teeth.
This form works by grasping food and grinding it before pumping the crushed material into the esophagus.
Malleate trophi are present in such common rotifers as
Keratella and Kellicottia. “Malleoramate” trophi (Fig.
8I) are found only in the order Flosculariacea and resemble the malleate form except that, in the malleoramate form, the rami are strongly toothed and the unci
possess many thin teeth. Similar to the malleoramate
form, “ramate” trophi (Fig. 9) have large, semicircular
shaped rami and unci with many teeth. Ramate trophi
are generally considered to be limited to the Bdelloidae
(Melone et al., 1998a). Only members of the order
Collothecacea (family Collothecidae) possess “uncinate” trophi (Fig. 8H). These trophi are characterized
by unci possessing few teeth, usually with one large
and a few small ones. “Virgate” trophi (Fig. 8D, K – O,
Q, S) are modified for piercing and pumping and generally can be recognized by the long fulcrum and
manubria and the presence of a powerful hypopharyngeal muscle. Some trophi of this form are asymmetrical. Virgate trophi are found in the common genera
Notommata, Polyarthra, and Synchaeta. “Forcipate”
trophi (Fig. 8J) as the name implies, have an action like
forceps, whereby the trophi are projected from the
mouth to grasp prey, which are then brought into the
mouth and swallowed. Forcipate trophi are limited
to the family Dicranophoridae. “Incudate” trophi
(Fig 8G, R) function by grasping prey with a forcepslike action, but this form has a different morphology
than the forcipate type; the rami are quite large and the
mallei very small. The mastax actually initiates prey
capture by creating a suction, drawing prey into the
mouth, which is then stuffed into the stomach with the
aid of the trophi. Asplanchna, which has no anus, uses
its trophi to extract undigestable materials from its
FIGURE 7 Generalized rotifer trophi. (Modified from Wallace
et al., 1989, with permission.)
202
Robert Lee Wallace and Terry W. Snell
FIGURE 8 Rotifer trophi types. (A) malleate trophi of Epiphanes, ventral; (B) malleate trophi (Epiphanes),
ventral elevated; (C) malleate trophi (Epiphanes), lateral; (D) virgate trophi of Notommata, dorsal; (E) malleate
trophi of Proales; (F) showing hypopharynx (lateral); (G) incudate trophi of Asplanchna; (H) uncinate trophi of
Collotheca; (I) malleoramate of Ptygura; (J) forcipate trophi of Dicranophorus; (K) asymmetrical virgate trophi
of Trichocera rattus; (L) virgate trophi of Eothinia; (M) virgate trophi of Itura, oral plates enlarged; (N) virgate
trophi of Synchaeta with the powerful hypopharynx muscle; (O) virgate trophi of Ascomorpha; (P) malleate
trophi of Proales gigantea, an intermediate form between malleate and virgate types; (Q) virgate trophi of
Cephalodella, dorsal; (R) incudate trophi of Asplanchna; and (S) virgate trophi of Cephalodella, lateral. Bars
20 m. (From Koste and Shiel, 1987, with permission.)
stomach. Incudate trophi are limited to the family Asplanchnidae. “Cardate” trophi (Fig. 10) are found only
in the family Lindiidae and function by producing a
pumping action, without the hypopharyngeal muscle.
The “fulcrate” type of trophi have been described as an
aberrant form and are incompletely understood (Edmondson, 1959, p. 432). This form is found only in the
class Seisonidea, a very small group of marine rotifers.
3. Organ Systems
Research on the ultrastructure of rotifers has proceeded rapidly during the past decade and promises to
continue to do so. For a comprehensive overview on the
topic of ultrastructural research on rotifers, consult the
works of Amsellem and Ricci (1982), Clément (1977,
1980, 1987), Clément et al. (1983), Clément and Wurdak (1991), Wurdak (1987), Wurdak and Gilbert
(1976), and Wurdak et al. (1977, 1983).
FIGURE 9 Ramate trophi of bdelloid rotifers. (Modified from
several sources.)
a. Muscular System The muscular system consists
of small groups of longitudinal and circular muscles inserted at various points on the integument or between
the integument and viscera (Fig. 2). In loricate species,
8. Phylum Rotifera
203
FIGURE 10 Cardate trophi of the family Lindiidae. (From Harring
and Myers, 1922, with permission.)
the integument provides a firm structure against which
the muscles work. Muscle contraction can increase the
pressure of the pseudocoel, which then acts as a hydrostatic skeleton. In some species, muscles retract the
corona, which increases pressure within the pseudocoel, thus expanding flaccid portions of the integument,
known as body-wall outgrowths (Asplanchna, Fig. 11).
In others, such as Brachionus (Fig. 12), retracting the
corona makes spines, that articulate with the body in
the posterolateral region, swing forward. When in this
extended position these spines interfer with predators
like Asplanchna which would otherwise consume the
Brachionus (Gilbert, 1966, 1967). Muscles also are
present in the viscera, particularly in the mastax and
stomach. Striated, longitudinal muscles are responsible
for retracting the corona and foot and for moving certain articulating spines that are not positioned by hydrostatic pressure. Some species possess powerful muscles that control movement of certain locomotory
appendages (e.g., Hexarthra, Fig. 13). Contractions of
these muscles cause a swift downward sweep of the
FIGURE 12
Spined and unspined Brachionus calyciflorus. (From
Koste, 1976, with permission.)
appendages which results in a rapid displacement or
jump of the rotifer (see Section II.C.1).
b. Nervous System The nervous system is simple,
consisting of only a cerebral ganglion or brain (Fig. 3)
located dorsally on the mastax, a few other ganglia present in the mastax and foot, and three types of sensory
organs, namely mechano-, chemo-, and photoreceptors.
Mechanoreceptive bristles are situated on the corona,
while several antennae are located elsewhere on the
body surface, usually laterally and caudally (Fig. 2).
Chemoreceptive pores are also present on the corona.
Many species possess one or more photoreceptive eyespots, sometimes accompanied by a pigmented spot
(Fig. 3). When present, eyespots are located in the anterior end, usually near the brain (Clément, 1980; Clément et al., 1983). While most rotifers retain the eyespots throughout life, larvae of sessile rotifers often lose
them during metamorphosis. Paired, ventral nerve cords
proceed from the brain along the length of the body
into the foot. Several other ganglia are usually found in
the nerve cords at the exit points for lateral nerves.
One interesting structure found in the apical region
of many bdelloid and monogonont rotifers is the
FIGURE 11
Body form variability in the genus Asplanchna.
(b.w.o. body-wall outgrowths). (Modified from Gilbert, 1980a,
with permission.)
FIGURE 13 Hexarthra showing positioning of muscles that initiate
jumps. (Modified from several sources.)
204
Robert Lee Wallace and Terry W. Snell
retrocerebral organ. This structure consists of paired
subcerebral glands and an unpaired retrocerebral sac,
both with ducts that lead to the surface of the corona
(Fig. 8.1). While the function of the retrocerebral organ
is unknown, Edmondson (1959, p. 422) has noted that
there are fewer protonephridia in rotifers that possess
well-developed retrocerebral organs. However, Clément
(1977) suggests that it may function as an exocrine
gland, perhaps lubricating the anterior part of the
body.
Although information on neurobiochemistry is
very limited, research has shown that acetylcholine
functions as a neurotransmitter (i.e., a cholinergic system) in twelve species of rotifers from six families (Nogrady and Alai, 1983, Nogrady and Keshmirian,
1986a, b). In addition, norepinephrine neuroreceptor
sites (i.e., an adrenergic system) have been reported in
Brachionus plicatilis, and a widespread catecholaminergic neuronal systems have been observed in species of
Asplanchna and Brachionus (see Keshmirian and Nogrady, 1988).
c. Protonephridium A paired protonephridial system comprised of tubules and flame cells functions in
excretion and osmoregulation in all rotifers (Figs. 1
and 2). Usually there are only a small number (6) of
flame cells, but large rotifers may possess many more.
For example, Asplanchna sieboldi may have up to 100
flame cells (Ruttner-Kolisko, 1974, p. 11). Normally,
the tubules drain into a urinary bladder which leads to
a cloaca, but the bladder is absent in some species and
a contractile cloaca assumes its function.
d. Reproductive System Besides their separation by
anatomical details of the trophi, the three classes of rotifers are differentiated based on the anatomy of their
reproductive systems (Wallace, 1998). The gonads are
paired in both the Seisonidea and the Bdelloidea, but in
the latter class, males are completely unknown and reproduction is always asexual. Members of the third
class (Monogononta) have only one gonad. Although
males are present in this group, they have not been described for a large number of species. However, it is
generally assumed that most (all) monogononts are capable of producing males given the proper conditions,
or at least that the ancestral forms were capable of
male production. When males are produced by monogonont populations, they are usually limited to a few
days or a week, so that during the year most reproduction is parthenogenetic (see Section III.B.1).
The reproductive organs of female rotifers comprise three units, which may be seen with a light microscope: ovary, vitellarium, and follicular layer (Amsellem and Ricci, 1982). The ovary of a rotifer is a
small, syncytial mass that is closely associated with the
yolk-producing vitellarium. At birth the adult complement of ovocytes already has been formed in the ovary.
The vitellarium is also syncytial with a constant number of nuclei, a characteristic useful in the taxonomy
of some species. The follicular layer surrounds both
the ovary and the vitellarium; in some species, this
layer forms the oviduct, which connects with the posterior portion of the gut forming a joint exit, the cloaca
(Fig. 1).
In general, monogonont males are much smaller
(about 100 m long) than females. In nearly all forms,
the gut of males is reduced to a rudiment or is absent
entirely (Fig. 2). In those forms with a rudimentary gut
(e.g., Asplanchna), it serves as an energy source for the
fast swimming, nonfeeding male. The single testis is
large and saccate, usually containing 50 freely floating mature sperm. A ciliated vas deferens leads from
the testis to the penis, usually with one or rarely two
pairs of accessory (prostate) glands that discharge into
it (see Section III.B.2).
Commonly, rotifers are oviparous, that is, they release their eggs outside the body where the embryos develop. Many planktonic rotifers carry their eggs attached to the body of the mother by a thin thread
(Brachionus), while others fix them to a substrate (Euchlanis) or release them into the plankton (Notholca).
A few species retain the embryo in the body until the
offspring hatches: ovoviviparous (e.g., Asplanchna and
Cupelopagis).
C. Environmental Physiology
1. Locomotion
All rotifers swim during at least a portion of their
life cycle and some swim continuously, never attaching
even temporarily to surfaces. Swimming influences the
acquistion of food and mates, increases the number of
encounters with predators, and promotes the dispersal
of the larvae of sessile forms. Some rotifers are unusual
in that they can swim, but routinely remain in mucus
tubes attached to a substrate (e. g., certain Cephalodella
species), while others are free-floating mucus sheaths
(Ascomorpha).
Most rotifers swim in a helical pattern, so that the
actual distance traveled is greater than the linear displacement (see Starkweather, 1987). However, for practical reasons, most researchers do not attempt to calculate absolute distance when considering the distance
traveled.
Although the theoretical power requirements of
swimming (i.e., the theoretical energy required to overcome water resistance) is 1% of the total metabolism,
the actual energetic cost appears to be much greater.
8. Phylum Rotifera
In Brachionus plicatilis, Epp and Lewis (1984) calculated this cost to be approximately 38% of the total
metabolism (See also Epp and Lewis, 1979).
Swimming speeds of male and female Brachionus
plicatilis have been measured by Luciani et al. (1983),
Epp and Lewis (1984), and Snell and Garman (1986).
Swimming speeds also have been measured for Asplanchna brightwelli (Coulon et al., 1983), two species
of Keratella (Gilbert and Kirk, 1988), and for Anuraeopsis fissa, Asplanchna girodi, Brachionus calyciflorus, Euchlanis dilatata, K. americana, Lecane quadridentata, Brachionus patulus, and Trichocerca pusilla
(Rico-Martinez and Snell, 1998). These studies have
shown that swimming speed is temperature – dependent
and varies among strains. Values recorded for B. plicatilis at 25°C range from 0.6 to 0.9 mm/sec for females
and from 1.3 to 1.5 mm/sec for males, with young and
old females swimming about 30% slower than mature
females. Asplanchna brightwelli and Keratella females
normally swim at 0.9 and 0.5 mm/sec, respectively. Euchlanis dilatata is one of the fastest species, with males
and females both swimming at 0.98 mm/sec at 25°C.
Little is known about the swimming speeds of the
larvae of sessile rotifers, but the swimming speeds of
the larvae of Ptygura beauchampi are know to vary
with age. New born (0 – 2 h) through mid-aged (ca., 3
h) larvae swam about 2 – 2.5 mm/sec, faster than the
rates reported for Brachionus males. However, older
larvae (4.5 h) swam at approximately 1.0 mm/sec.
Concomitant with these changes in swimming speed
was an increase in turning frequency (Wallace, 1980).
Correlations of rotifer ultrastructure with turning
frequency, locomotion in general, and other behaviors
have been a field of systematic investigation by Clément and his co-workers (Clément, 1987; Clément et
al., 1983; Chauoy, 1995). Their efforts have demonstrated that rotifers are excellent models for comparative neurobehavioral studies, but much more work remains to be done before a complete synthesis of
structure, function, and behavior will be possible.
Spines and other appendages influence both swimming speed and sinking rates in rotifers. For example,
Stemberger (1988) has found that unspined Keratella
testudo generally swim faster and sink more slowly
than spined forms. On the other hand, Polyarthra normally swims at a much slower velocity (0.24 mm/sec)
than any species previously discussed, but is capable of
very short bursts (about 0.065s long) of rapid movements called jumps. During a jump, Polyarthra may
attain velocities greater than 50 mm/sec, with a mean
velocity of 35 mm/sec, or 100 times its normal swimming speed (Gilbert, 1985a, 1987; Starkweather,
1987). Jumps are produced by movements of 12 appendages, called paddles, which articulate with the
205
body near the head. When Polyarthra detect a local
disturbance in the water a few body lengths away,
powerful striated muscles rapidly flex some of the paddles upward and then during the jump they return to
their original positions. During this process the rotifer
is displaced an average of 1.25 mm (ca. 12 body
lengths). Jumping helps this rotifer escape invertebrate
predators, such as Asplanchna (Gilbert, 1980b; Gilbert
and Williamson, 1978) and first instar Chaoborus
(Moore and Gilbert, 1987) as well as the filtering currents of microcrustaceans, such as Daphnia (Gilbert,
1985b, 1987). They are also effective against naive
workers attempting to remove Polyarthra from plankton samples using micropipets! The genera Filinia and
Hexarthra also can jump rapidly, but movement in
these forms has not been well studied (see Section
III.C.4). In contrast, Keratella cannot jump to avoid the
filtering currents of daphnids and, as a result, are severely damaged or killed when swept into the branchial
chambers of cladocerans (Burns and Gilbert, 1986a,b;
Gilbert and Stemberger, 1985b). However, Keratella is
not without some escape abilities; it is capable of increasing its swimming speed by a factor of about 3.5
when it encounters inhalant currents of Daphnia or the
predatory rotifer Asplanchna (Gilbert and Kirk, 1988).
Another source of phenotypic variation in body
size is polyploidy. Substantial intrapopulational variation in body size was reported for Euchlanis dilatata
(Walsh and Zhang, 1992). Two distinct body sizes were
observed in Devils Lake, Oregon over 12 months of
intensive sampling. The smaller morphotype was
about 225 m in length and the larger averaged about
275 m. The larger morphotype had a chromosome
number of 21 and the smaller morphotype had 14. The
haploid chromosome number found in males of this
species is n 7. This is the first case of polyploidy reported for rotifers, but new techniques for staining and
counting rotifer chromosomes may make more systematic explorations possible.
2. Physiological Ecology
Physiological tolerances of organisms prescribe environments where survival and reproduction are possible. Thus, an environmental tolerance curve for a
species summarizes the range of environments where
reproduction occurs and culminates at the upper and
lower lethal limits for the species (e.g., temperature
range). Within the tolerance curve, the environmental
optimum for a species is that environment where survival and reproduction are maximal. Therefore, a set
of tolerance curves (including temperature, pH, etc.)
indicate the breadth of adaptation of a species and its
niche width. Environmental tolerances and niche
widths have been characterized for a few rotifer
206
Robert Lee Wallace and Terry W. Snell
species. For example, Epp and Lewis (1980) described
the response of Brachionus plicatilis to temperature.
That study determined respiration rate over temperature ranging from 15 to 32°C and recorded Q10s of
1.9 – 2.4 for broad temperature intervals. Respiration
levels off between 20 and 28°C, indicating that B. plicatilis can maintain a constant metabolic rate over this
temperature range. At higher and lower temperatures,
respiration rate increased, presumably because of thermal stress which was beyond the homeostatic capability of this rotifer.
Snell (1986) showed that amictic and mictic females
have similar temperature tolerance curves. Amictic B.
plicatilis females reproduced at 20 and 40°C, whereas
mictic females did not. In general, amictic females reproduced over a broader environmental range of temperatures salinity, and food level than did mictic females.
The effect of pH on the distribution and abundance
of rotifers is a topic that has received a good deal of attention. However, since hydrogen-ion concentration is
related to other important chemical parameters in freshwaters, studies of rotifer occurrence as a function of pH
alone are of limited value. Nevertheless, some extensive
early work by Myers in the 1930s demonstrated that
rotifer species can be classified into a few broad groups
based on pH alone: alkaline species, acid species; and
those with a broad range (see also Edmondson, 1944).
More recently, Berzins and Pejler (1987) concluded that
species found in oligotrophic waters had pH optima at
or below neutrality (pH of 7.0) and those species common to eutrophic waters had optima at or above neutrality. Further, they noted that acid-water species were
often nonplanktonic or semiplanktonic.
Much less is known of the specific metabolic responses of rotifers to pH. The influence of pH on B. plicatilis was described by Epp and Winston (1978). They
found that swimming activity and respiration rate were
not significantly different at pH values of 6.5 – 8.5. Snell
et al. (1987) examined the pH from 4.0 to 9.9 and
found swimming activity depressed below pH 5.6 and
above pH 8.7. Alkaline waters depressed swimming activity more than acidic conditions. Increasing acidification of lakes as a result of acid rain has been widespread in North America. Acidification leads to
predicable changes in the rotifer zooplankton, such as
the increasing dominance of Keratella tauracephala
(Brett, 1989). This observation has been repeated in the
experimental acidification of lakes (Gonzalez and Frost,
1994). In general, it appears that the importance of
rotifers within lakes increases with acidity (Frost et al.,
1998).
Osmoregulation has been investigated by Epp and
Winston (1977) for B. plicatilis, a species that is common in highly alkaline waters and salt lakes. This species
was an osmoconformer capable of tolerating osmolarities exceeding 957 mOsm/L (33.5 ppt). Ito (1960)
suggested that the upper osmotic limit for B. plicatilis
may be as high as 2860 mOsm/L (97 ppt). Transfer of
this rotifer directly from 41 to 957 mOsm/L caused considerable mortality, but acclimation to high osmolarities
was achieved by gradually increasing ionic concentrations. Brachionus plicatilis does not tolerate low
osmolarities well, and this probably accounts for its restriction to alkaline and brackish waters.
Although most rotifers require oxygen concentrations significantly above 1.0 mg/L, some can tolerate
anaerobic or near-anaerobic conditions for short periods. Other species routinely live in oxygen-poor regions, such as the hypolimnion of eutrophic lakes or in
sewage ponds. The physical and chemical factors described above, along with food (Bogdan and Gilbert,
1987) and predation (Williamson, 1983), define the
niche boundaries for rotifer species. Niche boundaries
for field populations have been described by Edmondson (1944), Makarewicz and Likens (1975, 1979),
Miracle (1974), and Miracle et al. (1987). These investigations have generally revealed that rotifer species
extensively partition the environment, thus avoiding
negative interactions.
3. Environmental Toxicology
Because rotifers fill an important ecological role
and are relatively easy to culture, interest is growing in
their use for aquatic toxicity testing. The response of
rotifers to a variety of toxicants has been characterized
in both natural and laboratory populations. For example, effects of various insecticides, herbicides, and
wastewater on natural rotifer populations have been
investigated (Snell and Janssen, 1995). In general, rotifers seem to serve as good indicators of environmental
water quality (Sladecek, 1983), and the use of rotifer
population dynamics as sensitive indicators of toxicity
has been promoted (Halbach, 1984; Halbach et al.,
1981, 1983). A multispecies approach to toxicity assessment with a laboratory microcosm consisting primarily of rotifers has been attempted by Jenkins and
Buikema (1985). Many short-term, acute toxicity tests
with a variety of substances have been conducted using
rotifers, e.g., insecticides, heavy metals, free ammonia,
sodium dodecyl sulfate, crude oil and petrochemicals
(Snell and Janssen, 1998).
Unfortunately, the results of these studies are not
directly comparable, as their methodologies varied
considerably. However, testing protocols are comparable from several studies of the halophile B. plicatilis
and a number of median lethal concentrations have
been reported. (LC50 is the concentration of a compound which kills 50% of a cohort within a specified
8. Phylum Rotifera
exposure time.) Capuzzo (1979a, b) found B. plicatilis
to be very sensitive to free chlorine and chloramine. After a 30-min exposure at 25°C, LC50 values of 0.09 and
0.01 mg/L were recorded for chlorine and chloramine, respectively. Brachionus is more sensitive to
mercury (LC50 of 0.045 mg/L; Gvozdov, 1986) than to
chromium (LC50 500 mg/L; Persoone et al., 1989).
Free ammonia is also toxic, with LC50 values of 20.4
and 17.7 mg/L at salinities of 15 and 30 ppt, respectively (Snell and Janssen, 1995).
Pesticides, in contrast, are not very toxic to B. plicatilis. The 24-h LC50 values for five organophosphate
pesticides and one organochlorine pesticide ranged
from 0.9 to 150 mg/L for three different strains of B.
plicatilis (Serrano et al., 1986). These same investigators observed that pesticide resistance of this rotifer is
about 1000 times greater than that reported for other
aquatic organisms. Since pesticides are generally targeted to arthropods, the low sensitivity of rotifers to
these compounds is not surprising.
Several median lethal concentrations for a variety of
compounds have been reported for freshwater rotifers,
mainly in the genus Brachionus. Dad and Kant Pandya
(1982) recorded 24-h LC50 values for B. calyciflorus exposed to two insecticides, and Couillard et al. (1989)
found the following relative metal toxicities: Hg Cu
Cd Zn Fe Mn. LC50 values of 600 and 0.16
mg/L for phenol and pentachlorophenol, respectively,
were reported by Halbach et al. (1983) for B. rubens.
A standardized rotifer toxicity test for marine and
freshwater that employs test animals derived from
hatching B. plicatilis or B. calyciflorus resting eggs has
been described (ASTM, 1991). This test is less expensive
because stock cultures are not necessary to obtain test
animals, and it is simple, rapid, and sensitive, so the use
of rotifers in aquatic toxicology is expected to expand.
4. Anhydrobiosis
The ability of rotifers to tolerate desiccation and
then be revived sometime later has been known since
the early 1700s, when Leeuwenhoek observed rehydration of rotifers found in dry sediments of rain gutters.
Anhydrobiosis, which is also termed cryptobiosis and
osmobiosis, is limited to bdelloids. Unlike monogononts, bdelloids cannot produce cysts. The term
cryptobiosis refers to the slow or hidden metabolism
which is characteristic of these animals, while anhydrobiosis and osmobiosis emphasize the processes whereby
loss of water is accomplished through evaporation and
external osmotic pressure, respectively. While in the
desiccated state, these rotifers resemble a wrinkled barrel (or tun, as in the tardigrades, see Chapter 15), with
the head and foot retracted into the animal’s trunk
(Fig. 14). The significance of this phenomenon is clear,
207
FIGURE 14
Body of a desiccated bdelloid rotifer (Habrotrocha
rosa) with the corona withdrawn into the trunk. (Bar 50 m.)
as many bdelloids inhabit environments that dry completely at irregular intervals. A few strictly aquatic
bdelloids cannot withstand desiccation (Ricci, 1998b).
Anhydrobiosis involves more than simple drying;
unless loss of metabolic water proceeds slowly, the rotifer usually dies. During anhydrobiosis, the fine structure of cells is retained, but in a greatly modified state
(Ricci, 1987; Schramm and Becker, 1987). Changes
that occur internally include a 50% reduction in the
volume of the pseudocoel, a condensation of cells and
organs, and a decrease in cytoplasmic volume, so that
the entire animal is only about 25 – 30% of its original
size. Nuclei, mitochondria, endoplasmic reticula, and
other organelles form a compact mass within cells of
the anhydrobiotic animal (Schramm and Becker, 1987).
Desiccated bdelloids have been reported to be viable even after more than two decades in the anhydrobiotic state. Recovery from anhydrobiosis may require
as little as 10 min, or it may take several hours, according to prevailing environmental conditions. Survival is
negatively affected by starvation before desiccation and
by moist environments and high temperatures during
the desiccation period (Ricci 1987; Ricci, et al., 1987,
Schramm and Becker, 1987).
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
1. Phenotypic Variation
Phenotypic variation is an important adaptive
mechanism in rotifers, but has posed difficult problems
for systematists. This variation arises by several mechanisms including cyclomorphosis, dietary- and predatorinduced polymorphisms, polymorphisms in hatchlings
208
Robert Lee Wallace and Terry W. Snell
from resting eggs, and dwarfism. Cyclomorphosis is the
seasonal phenotypic change in body size, spine length,
pigmentation, or ornamentation found in successive
generations of zooplankton. These changes are phenotypic alterations in a single population that are related
to physical, chemical, or biologic features of the environment. Each different morphological form is called a
morphotype. Specifically excluded from cyclomorphotic
change are seasonal succession of sibling species and
clonal replacements of genotypes, both of which are genetic changes in populations.
A striking phenotypic change in morphology that
is associated with a dietary polymorphism was described for three Asplanchna species (brightwelli, intermedia, sieboldi) by Gilbert (1980a). Diets that include
the plant product a-tocopherol (vitamin E) induce saccate females, the smallest morphotype, to produce cruciform daughters. Cruciforms have lateral outgrowths
of the body wall (Fig. 11) that protect them from cannibalism by conspecifics by making them larger and,
thus, more difficult to ingest if captured. In the presence of a-tocopherol and certain prey types, cruciforms
can produce a third morphotype called campanulates
(more prevalent in A. sieboldi and A. intermedia).
Campanulates are very large females ( 2000 m),
which heavily cannibalize saccate females. Female polymorphism is much less pronounced in A. brightwelli
where there is a 50 – 60% increase in body size, but no
campanulates are produced and body wall outgrowths
are slight. Dietary polymorphism in Asplanchna (gigantism) may have evolved originally as a generalized
growth response to larger prey typical of eutrophic waters (Gilbert, 1980a; Gilbert and Stemberger, 1985c).
The tocopherol response probably is adaptive, because
it signals the availability of nutritious rotifer and microcrustacean prey.
Another source of phenotypic variation is predator-induced polymorphisms. Spined and unspined
forms had been recognized in several rotifer species for
many years, but the cause(s) and significance(s) of these
variations remained an enigma (Fig. 12). However,
Gilbert (1966, 1967) was the first to show that spine
production could be induced in the offspring of female
B. calyciflorus if adults were exposed to culture
medium which had previously held the predatory
species Asplanchna. Gilbert (1967) also demonstrated
that such spines were strong deterrents to predation by
Asplanchna (see Section III.C.4). However, Stemberger
(1990) has shown that food concentration can dramatically modify the development of spines in B. calyciflorus.
Two additional sources of phenotypic variation are
polymorphisms called “aptera generations” in the
hatchlings of resting eggs and dwarfism, both of which
have been reported in rotifers of some tropical crater
lakes (Green, 1977). Aptera morphotypes were initially
thought to be different species of Polyarthra, but later
were shown to be forms lacking the paddles that
are characteristic of this genus. Only the generation
hatching from resting eggs lacks paddles; their
parthenogenetic offspring develop into typical morphotypes. Similar polymorphisms between resting egg
hatchlings and parthenogenetic generations were described for Keratella quadrata and are suspected for
Notholca acuminata (Amrén, 1964). Dwarfism in Brachionus caudatus in Cameroon crater lakes was
described by Green (1977) and is characterized by
reduced body size and spination as compared to normal morphotypes. Green speculated that high temperature combined with reduced food supply may cause
this condition.
2. Distribution and Population Movements
Water bodies are not uniform habitats with respect
to concentrations of food and predators and abiotic
factors such as dissolved oxygen concentration, light
intensity, temperature, and water movements. Therefore, it is not surprising to find that rotifers are not
evenly distributed in lakes and ponds; often there is
considerable variability with respect to their horizontal
and vertical distribution. For example, Hoffman (1982)
showed that two Filinia species had very different vertical distribution patterns over the course of a year in
a small lake in Germany (Fig. 15). Striking horizontal
variabilty in rotifer distribution has been reported by a
variety of researchers: for example, by Green (1985)
for a tropical lake. Thus, we find that some rotifers are
strictly littoral, being found in open waters only as occasional migrants, whereas others are pelagic; however,
the depth at which they are found is a function of season. Consequently, experienced researchers can determine to some degree where and when a water sample
has been taken, merely by the composition of the rotifer species it contains. Although these major distribution patterns are the result of differential population
growth and water currents and other large-scale water
movements, within-lake distribution patterns can be influenced to a lesser degree by locomotory behaviors.
One commonly recognized behavior of marine and
freshwater zooplankton is a daily (diel) vertical migration in the water column in which the animals usually
come to the surface only during the night. During the
day, zooplankton avoid visual predators (fish) that occupy near-surface waters, but when they return to the
surface at night they can exploit the rich algal resources
present there. In rotifers, diel migrations are never as
dramatic as those typical of microcrustaceans; the population maximum usually changes only about 1 – 2 m
8. Phylum Rotifera
209
FIGURE 15 Temporal and spatial distribution of two species of Filinia in Lake Plußee, Germany. Solid line
indicates limits of the population at one individual per liter; dotted line indicated boundaries of higher population
levels (numbers of individuals per liter). (Modified from data in Hofmann, 1982, with permission.)
over a daily cycle (Fig. 16). However, ovigerous (eggbearing) and nonovigerous females may have different
migration patterns, and this can cause serious errors in
the calculations of birth rates. Errors of nearly an order
of magnitude depending on the sampling protocol may
occur, if the population is sampled at only one depth or
at different times during the day (Magnien and Gilbert,
1983).
Horizontal movement of rotifers has been investigated by Preissler (1980) who showed that some
pelagic rotifers apparently avoid the inshore region.
Preissler demonstrated this phenomenon, known as
“avoidance of the shore,” by using a circular plexiglass
arena in which he simultaneously monitored the swimming direction of several zooplankton species. When he
artifically altered the shadow produced by the natural
elevation of the shoreline by adding a black collar
around the arena, both Asplanchna priodonta and Syn-
chaeta pectinata swam away from the shadow. The littoral rotifer Euchlanis dilatata apparently showed no
preference based on light intensity.
Research on the abundance and distribution of rotifers in lotic systems (flowing waters) has lagged far
behind that of lentic ones, but we do know that both
rotifer diversity and abundance are usually much lower
in rivers than in lakes. Further, most of what is present
seems to be derived from upstream lakes or secondary
channels with minimal reproduction within the channel
(Saunders and Lewis, 1989). In some instances, the
number of species is very high, but usually only a few
of these are numerically significant (10). The density
of rotifers encountered in lotic (flowing) systems is
usually much smaller than that commonly present in
lentic (lakes) ones (ca. 5 – 400 individuals/L), but in
some instances it may be quite high (ca. 3,000 individuals/L).
210
Robert Lee Wallace and Terry W. Snell
FIGURE 16
Depth distribution of Keratella crassa in a small, shallow lake during a 48 h period in the
summer of 1980. (A) Total population; (B) nonovigerous population; (C) ovigerous population; (D) egg ratios
calculated for each sampling point; (E) egg ratio and mean density of the entire population throughout the
water column. Kite diagrams express the population as relative percentages for each sampling point, while the
line indicates the mean depth of the entire population. (From Magnien and Gilbert, 1983, with permission.)
3. Biogeography
Early studies of rotifer distributions were dominated by the belief that all rotifers are cosmopolitan, an
idea supported by the fact that the resting eggs of
monogononts and the anhydrobiotic stages of bdelloids
are transported by birds and other mechanisms. Such
passive dispersal, it was argued, is very effective at ensuring that most species become globally distributed.
However, data has accumulated suggesting that this
conclusion was premature. Rotifers may not be as easily dispersed as previously thought (Jenkins and Underwood, 1998). Green (1972) showed a latitudinal zonation in certain planktonic rotifers, and Pejler (1977b)
provided data demonstrating that some species have restricted distributions. In general, endemism seems to be
an important theme in several genera, including Keratella, Notholca, and Synchaeta (Dumont, 1983). Exhaustive inventories of rotifer fauna in a few lakes has
suggested that 150 – 160 rotifer species reside in a typical temperate lake as compared to more than 210
species for a tropical lake (Dumont and Segers, 1996).
These numbers are likely to go higher as sessile and
contractile species become more well known. These authors argue that the theory of cosmopolitanism must be
abandoned for rotifers and cladocerans.
In comparison to the substantial research effort
developed elsewhere, there has been relatively little
work on rotifer biogeography in the United States and
most of that work was done prior to the 1950s or
is limited to specific geographic areas (e.g., the Great
Lakes). Nevertheless, in North America there are
several examples of species with restricted distributions
in the genera Anuraeopsis, Brachionus, Lecane, and
Lepadella. Central America and the southern United
States have close affinities to tropical South American
fauna, while further north, affinities with Europe are
apparent and endemism is strong in Keratella,
Notholca, and Synchaeta (Chengalath and Koste,
1987). Dumont (1983) suggested that continental drift
and Pleistocene glaciations best explain the current biogeography of rotifers.
Most biogeographical analyses of rotifers have
been limited to collating information on the location of
populations from descriptions found in the literature.
This problem has additional difficulty because many
species undergo periodic polymorphism (cyclomorphosis) (see Section III.A.1), with each population having a
slightly different morphology. In the past, this phenomenon confused some workers who considered each
morphotype to be a new species. Furthermore, the
question of what constitutes a rotifer species becomes
problematic as males have never been described for
many taxa. Future efforts to clarify these matters probably will be greatly aided by modern techniques in gel
electrophoresis (e.g., King, 1977; King and Zhao,
1987; Snell and Winkler, 1984) and by using mating
behavior to define species boundaries (Snell and
Hawkinson, 1983; Snell et al., 1988; Snell, 1989).
In the future, biogeographic research may play an
important role in monitoring environmental conditions
by examining the structure of rotifer communities in a
variety of lakes subject to environmental pertubations:
8. Phylum Rotifera
e.g., heavy metal input (MacIsaac et al., 1987) and acid
precipitation (Frost et al., 1998).
4. Colonial Rotifers
Most rotifers are solitary and interact only as potential prey or mates, but about 25 species (in eight
genera) of the class Monogononta form permanent
colonies (Wallace, 1987). All colonial forms are members of two families (Flosculariidae and Conochilidae)
of the order Flosculariacea (Fig. 17). None of these
taxa are predators and all reproduce in a fashion typical of monogononts. There appears to be a relationship
between coloniality and sessile existence. About 70%
of colonial forms (18 species) are sessile, but even more
211
striking is the fact that all seven genera of the family
Flosculariidae have colonial species. This has led some
workers to suggest that sessile species are somehow
preadapted for the evolution of coloniality (see Wallace, 1987 for a review). The form of colony production has significant implications. Because colonial rotifers do not reproduce by budding or the formation of
specialized zooids, colony members are not intimately
connected, as are the colonial bryozoans (Chapter 14);
therefore, energy resources cannot be shared among
colony members. Some colonial rotifers produce tubes
from either hardened secretions (e.g., Limnias), pellets
(e.g., Floscularia conifera), or gelatinous secretions
(e.g., Lacinularia and Conochilus). These tubes are
FIGURE 17 Colonial rotifers. (A) A portion of a small, planktonic colony of Conochilus unicornis: animals
live within a gelatinous mass which they secrete, also present are many small euglenoids; (B) a small, sessile
colony of Floscularia: adults reside within tubes they construct of tiny pellets of compacted detritus; and (C) a
large sessile colony of Sinantherina.
212
Robert Lee Wallace and Terry W. Snell
important in the overall structure of the colony as they
provide a substrate or matrix for the addition of new
members to the colony (see Section III.A.5).
The number of individuals in a colony varies
greatly among genera. Floscularia ringens usually
builds colonies with fewer than five individuals, as do
some members of the family Conochilidae. In this family, small colonies, composed of only an adult and oneto-several young, are produced by some species, while
others may have more than 250 individuals within
a colony. Although most colonial species construct
small colonies of fewer than 5 – 24 individuals, some
routinely produce large colonies of 50 – 200 individuals
(e.g., Floscularia conifera, Sinantherina socialis,
Conochilus hippocrepis), and a few produce colonies
of truly gargantuan size (1000; Lacinularia).
Colonies form by one of three different methods,
but these methods produce colonies that may have a different degree of genetic relatedness among colony mates
(Table I). In allorecruitive (Greek, allo, “other”) colony
formation, free-swimming young (larvae) produce
colonies by settling on tubes of sessile adults. Thus, the
colonies of allorecruitive species grow in size (numbers
of individuals per colony) and in the number of colonies
within the habitat (colony density) by intercolonial recruitment of young. Because the larvae joining these
colonies may come from females belonging to another
colony, genotypic relatedness within the colony is probably low. These colonies are transitory, beginning when
larvae attach to a previously settled adult and ending
when larval recruitment to an old colony ceases. Late recruits to an allorecruitive colony may suffer reduced fecundity, because death of the founding individual might
lead to dislodgment from the substrate and death of all
members of the colony. A few allorecruitive species produce interspecific colonies of two or more species. Some
species of the genus Floscularia reproduce this way.
TABLE I
In autorecruitive (Greek, auto, “self”) colony formation, the young remain with their mother in the
colony. Thus, the colonies of autorecruitive species
grow in size (numbers of individuals per colony) by intracolonial recruitment of young back into the parental
colony and in the number of colonies within the habitat (colony density) by colony fission. Because the
young joining these colonies come from the parent
colony, genotypic relatedness within the colony is probably high. Here autorecruitive colonies are long-lived
and develop continuously throughout the season, increasing in size as new individuals are added and decreasing only if the colony divides. Late recruits to
these colonies probably do not suffer diminished fecundity because the colony continues though individuals
die. Autorecruitive species rarely produce interspecific
colonies. Species of the genera Conochilus and
Conochiloides reproduce this way.
In geminative (Latin, gemin, “double”) colony formation, all the young born within a span of a few
hours leave the parent colony as a free-swimming
larval colony. Members of this young colony subsequently explore and attach to a new substrate in
concert, but they may join an older colony, thus genotypic relatedness within the colony is probably moderate to high depending on the particular history of
the colony. Thus, because the size of the larval colonies
depends mainly on the fecundity of the parental colony,
the number of individuals in a colony does not increase
over its life time. Geminative species rarely produce
interspecific colonies, but it is possible. Species of
the genera Lacinularia and Sinantherina reproduce
this way.
The consequences of high genetic relatedness
have not been explored, but may include increased
vulnerability to parasites, uniformity of behaviors, and
decreased genetic diversity of resting eggs.
Colony Formation in Rotifers
Colony formation
category
Increase in colony
size (individuals/colony)
Allorecruitive
(intercolonial)
Autorecruitive
(intracolonial)
Young are recruited
to established
colonies or remain
solitary
Young stay within
the parental colony
Geminative
Rare-to-absent
Mode of increase of
colony density in the
habitat
Predicted
genotypic relatedness
within colonies
Frequency of
interspecific colony
formation
Young leave the
parental colony
and may establish
a new colony
Fission: adult
colonies fragment
producing smaller
colonies
Cohorts of larvae
collectively
establish new
colonies
Low
Common
Floscularia
(Fig. 17B
& 43A)
High
Absent
Conochilus
(Fig. 17A)
Moderate-to-high
Rare-to-absent
Lacinularia
(Fig. 5C)
Sinantherina
(Figs. 17C)
Examples
8. Phylum Rotifera
Two hypotheses have been offered to explain the
adaptive significance of coloniality in rotifers. One hypothesis suggests that colonial animals possess an energetic advantage over solitary individuals of the same
species; for example, colonial F. conifera apparently live
longer and mature faster than the solitary individuals
(Edmondson, 1945). Juxtaposition of filtering currents
produced by two individuals may permit increased filtering rate and/or enhanced filtering efficiency. Although
experiments have not supported the idea that coloniality
affects filtration rates, Wallace (1987) has provided
some information that supports the view that coloniality
increases filtering efficiency. A second hypothesis argues
that colonial existence can protect individuals from certain predators. Gilbert (1980b) and Diéguez and Balseiro (1998) showed that large Conochilus colonies are
less vulnerable to attack by certain invertebrate predators, because they are too large to be engulfed whole and
because individual rotifers can retract into the refuge of
the gelatinous matrix of the colony.
5. Sessile Rotifers
Sessile species are found in two families of rotifers:
Flosculariidae (7 genera) and Collothecidae (5 genera).
Although they are often overlooked because their habitats generally are not examined thoroughly, these forms
are actually quite common in lakes and rivers in North
America, occasionally reaching very high densities on
plant surfaces (6 individuals/mm2) especially in bogs
and small eutrophic ponds (Wallace, 1980; Wallace and
Edmondson, 1986).
FIGURE 18
213
The juvenile motile stages of sessile rotifers (Fig. 5)
are not considered to be larvae by some researchers, but
others have used this term and there is a conceptual parallel to other sessile invertebrates that undergo an extensive metamorphosis after settlement (Wallace, 1980).
Not surprisingly, the behavior of larvae changes dramatically once they come into contact with a potential attachment site. These new behaviors have been described
using terms such as selection, choice, and preference.
However, it should be recognized that the use of such
words does not imply cognition by the rotifer; they are
merely convenient terms to describe this phenomenon.
Several workers have demonstrated that larvae can
select a particular substrate from among those available for settlement (Wallace, 1980). For example, larvae of F. conifera settle with a greater frequency on the
tubes of conspecifics than on aquatic plants, although
there is substantially more plant surface available. This
propensity leads to the formation of intraspecific
colonies, each with 50 or more individuals. During the
growing season, about 75% of the entire population
may be colonial (Wallace, 1977).
Some species apparently attach to a surface based
on the chemistry of the water in the vicinity of the substrate. Wallace and Edmondson (1986) have shown
that greater than 90% of Collotheca gracilipes larvae
prefer the undersurface of Elodea canadensis leaves to
the upper surface (Fig. 18). Choice of the undersurface
is apparently in response to the way Elodea can alter
the concentration of calcium ions in the water immediately around the leaf. In water having a pH in the
Collotheca gracilipes colonizing the apical meristern of the aquatic
macrophyte Elodea canadensis. Many (25) individuals may be seen attached to the under
surfaces of the leaves. (Length of the lowest leaf is ⬇ 1 cm.) (From Wallace and Edmondson,
1986, with permission.)
214
Robert Lee Wallace and Terry W. Snell
neutral-to-alkaline range, Elodea acts as a polar plant,
removing Ca2 from beneath the leaf and releasing it
above. This choice provides a superior habitat in comparison to the upper surface. In short-term, laboratory
experiments, young C. gracilipes that attached to undersurfaces of Elodea leaves grew significantly taller
and produced more eggs per female than those attached to the upper surfaces of the same leaves. Several
other species of sessile rotifers exhibit strong preferences for particular substrates, but the significance of
these associations has not been fully elucidated (Wallace, 1980).
Larval substrate selection behaviors have been described for three species: Ptygura beauchampi, Sinantherina socialis, and C. gracilipes. In general, larvae appear to react to potential surfaces in similar ways. Very
young larvae avoid settling for periods up to several
hours after hatching. Once this refractory period is
past, all surfaces are explored, but some (i.e., the preferred substrates) receive much more attention and
elicit different behaviors, including some reminiscent of
male mating behavior (see Section III.B.2). A larva will
traverse these surfaces with both its corona and foot in
contact with the surface, occasionally stopping in this
slightly bent position. The larva may continue exploration of the surface for several minutes, but eventually
it attaches to the surface using cement from glands in
the foot and then undergoes metamorphosis.
Immediately after attachment and metamorphosis
have occurred, the young of most sessile rotifers begin
to secrete a protective tube. Often this secretion is in the
form of a clear, gelatinous material (e.g., Collotheca and
Stephanoceros), but in others the tube becomes somewhat opaque by the adherence of debris and colonizing
microorganisms (e.g., several Ptygura species and Lacinularia). One species of Limnias forms a cement tube
that looks like a series of rings, placed one on top of
another. Perhaps the most fascinating example of tube
construction is found in the genus Floscularia. Some
species in this genus possess a small ciliated cup on the
ventral side of the head to which some tiny mineral particles and other small debris collected by the corona are
shunted. The ciliated cup appears to be in constant motion, mixing gelatinous secretions with the particles to
form small pellets, either in the shape of bullets or balls.
Once a pellet is fully formed, the rotifer places it on top
of the tube in an action resembling the movements of a
bricklayer. In this way the tube is constantly elongated
as the animal grows. Because pellets are manufactured
from particles collected from the water, they may have a
greenish tint, but they are usually brown. When a heavy
rainstorm temporarily suspends soil in the water, the
pellets that are produced by Floscularia usually turn out
to be very dark. Thus, the event of the storm is marked
as a dark ring in the tubes of all the animals alive at
that time. Edmondson (1945) noted this and used suspensions of powdered carmine and carbon black to
mark the tubes of F. conifera, so that he could study the
dynamics of population growth and various aspects of
rotifer life history in a field population (See Fig. 17B).
B. Reproduction and Life History
1. Reproduction
The type of reproduction varies among the three
classes of rotifers (Wallace, 1998). Species in the class
Seisonidea reproduce exclusively through bisexual
means, with gametogenesis occurring via classical meiosis and the production of two polar bodies. At the other
extreme, members of the class Bdelloidea reproduce entirely by asexual parthenogenesis, a process which includes two equational divisions producing two polar
bodies. No males have been observed in bdelloids.
Species in the class Monogononta exhibit cyclical
parthenogenesis where asexual reproduction predominates, but sexual reproduction also occurs occasionally.
a. Cyclical Parthenogenesis Cyclical parthenogenesis in monogononts (Fig. 19) takes place in the absence
of males (amictic phase), but periodically males are
FIGURE 19
Generalized life cycle for monogonont rotifers. The life
cycle of bdelloids consists of only the parthenogenetic portion.
8. Phylum Rotifera
produced and sexual reproduction takes place (mictic
phase). Amictic females are diploid and produce
diploid eggs (called amictic eggs) that develop mitotically via a single equational division into females (Wallace, 1998). The term “summer egg” has been applied
to the eggs produced by amictic females, but this term
is misleading because amictic eggs can be produced
at any time of the year, depending on the species in
question.
Most of the monogonont rotifer life cycle is spent
in the amictic phase, but in certain conditions, sexual
reproduction occurs concurrently within the population, presumably initiated by specific environmental
stimuli. Upon receiving the mictic stimulus, amictic females begin producing both mictic and amictic daughters. The proportion of mictic daughters and the duration of their production, depends on the strength of the
mictic stimulus. Mictic female production is only the
initial step in mictic reproduction and is followed by
male production, fertilization, and finally formation of
a diapausing embryo called a resting egg or cyst (also
winter or fertilized egg) (Fig. 19). Mictic females produce haploid eggs via meiosis that, if unfertilized, develop into haploid males. Males are typically smaller,
faster swimming, and shorter lived than females. The
males of some species are structually reduced, but extremes exist from fully developed males (Rhinoglena)
to a condition in which certain organs are degenerate
(Conochilus). Among sessile species the male is freeswimming while the female is permanently attached to
a substrate.
Fertilized mictic eggs are diploid and develop into
resting eggs possessing thick, often sculptured, walls
(Fig. 20). These dormant stages are very resistant to
harsh environmental conditions and may be dispersed
over wide areas by the wind, water, or migrating animals. After a period of dormancy, which varies among
species, resting eggs respond to species-specific environmental cues and hatch, releasing diploid, amictic females that enter into the asexual phase of the life cycle
(Fig. 19). The stimuli that induce hatching may include
changes in light, temperature, salinity, and oxygen concentration (Wallace, 1998).
With a few notable exceptions, the stimulus for initiating sexual reproduction is still poorly understood.
Dietary -tocopherol (vitamin E) controls the shift
from amictic to mictic reproduction in most Asplanchna species (Gilbert, 1980a), and photoperiod
plays a similar regulatory role in Notommata (Pourriot
and Clément, 1981). In the genus Brachionus, population density has been most frequently cited as a stimulus for mictic female production (Wallace, 1998; cf,
Pourriot et al., 1986). In addition to environmental factors, genetic factors also play a major role in determin-
215
ing the sensitivity of particular strains to mictic stimuli
(e.g., Snell and Hoff, 1985; Lubzens et al., 1985).
An unusual variation of the standard monogonont
life cycle has been observed in three genera: Asplanchna, Conochilus, and Sinantherina. In some populations amphoteric females (individuals that produce
both female and male offspring) have been recorded
(e.g., Ruttner-Kolisko, 1977b; King and Snell, 1977).
Amphoteric rotifers have the ability to produce both
diploid (female) and haploid (male) eggs, with some
producing both females and resting eggs, or males and
resting eggs (Fig. 17). The presence of amphoteric reproduction in other rotifers has not been fully investigated, and its significance in the life history of those
genera that possess it remains to be determined. There
have been reports of the production of pseudosexual
eggs; that is, the production of resting eggs via
parthenogenesis in the absence of males. However, little
in known of this phenomenon.
Two different types of parthenogenetic amictic eggs
have been described in Synchaeta pectinata (Gilbert,
1995). The traditional type is thin shelled (1.4 m) that
develops without diapause. The second type has a
thick-shelled (9 m) amictic egg that enters an obligatory diapause after 1 – 3 cleavage divisions. The mean
duration of diapause is about 14 days, and it is not
greatly extended by low temperature. These eggs are
induced immediately after starvation and seem to be
produced at no additional energetic cost compared to
traditional amictic eggs. The adaptive significance of
diapausing amicitic eggs seems to be to increase the
ability of populations to survive short-term food limitation. Synchaeta pectinata also produce resting eggs following sexual reproduction and these have the capacity
for dormancy extending up to several years.
b. Resting Eggs The density of resting eggs in sediments has not been routinely examined and the few
published studies have used different methods for estimating density. Because of their capacity for extended
dormancy, resting eggs theoretically could accumulate
to high densities in sediments. Nipkow (1961) found
resting egg densities for seven species to range from 100
to 600 eggs/cm2 in lake sediments, but Synchaeta oblonga densities ranged from 3,000 to 4,000 eggs/cm2.
Snell et al., (1983) found an average of 200 resting
eggs/cm2 for Brachionus plicatilis in the surface sediments of a subtropical brackish pond. These workers
showed that the highest densities were on the sediment
surface, and numbers declined exponentially to a depth
of 7 cm. May (1987) examined resting egg densities in
sediments from Loch Leven, Scotland, by recording the
number of animals hatching from sediments incubated
at temperatures of 5, 10, and 15°C. This technique is
216
Robert Lee Wallace and Terry W. Snell
FIGURE 20
Resting eggs of several monogonont rotifers. (A) Asplanchna girodi; (B) Hexarthra fennica; (C)
Brachionus calyciflorus; (D) Conochiloides natans; (E) Sinantherina socialis; and (F) Kellicottia bostoniensis.
(Bars 10 m; the wrinkled backgrounds in panels B and F are the supporting membranes.) (Original scanning electron photomicrographs by Hendrik Segers.)
not directly comparable to the others just noted, and
probably yields lower estimates of resting egg density
(see also May, 1986). Marcus et al. (1994) observed
B. plicatilis resting eggs at densities up to 1532 individuals/cm2 in anoxic and sulfidic sediments of Pettaquamscutt estuary in Rhode Island. Rotifer resting eggs were
abundant as deep as 12 cm into these sediments and
were the most abundant among any of the zooplankton
taxa present. B. plicatilis resting eggs older than
40 years could be hatched and used to initiate viable
populations.
The number of resting eggs in sediments is a function of previous resting egg production, mortality in the
sediments, and hatching rates. Certain abiotic features
8. Phylum Rotifera
217
such as sedimentation rates and the uneven deposition
of sediments in the benthos (sediment focusing) may
affect the final density at particular sites in a lake or
estuary.
2. Reproductive Behavior
Detailed descriptions of mating behaviors are available for Asplanchna brightwelli (Aloia and Moretti,
1973), three species of Brachionus (e.g., Gilbert, 1963b;
Snell and Hawkinson, 1983), and Anuraeopsis fissa,
Asplanchna girodi, Euchlanis dilatata, Keratella americana, Lecane quadridentata, Brachionus patulus, and
Trichocerca pusilla (Rico-Martinez and Snell, 1998).
Males and females show pronounced sexual dimorphism, males being smaller and faster swimmers
(Fig. 2). Lacking a functional foot, males swim constantly without attaching. Because males and females
swim randomly, the probability of male – female encounters in planktonic species can be modeled mathematically (Snell and Garman, 1986). In the colonial,
sessile rotifer Sinantherina socialis, males may copulate
with several females of one colony in succession. Females take no active role in locating a mate or reacting
to the male once an encounter occurs. Males, in contrast, display a distinct mating behavior upon encountering conspecific females.
Mating behavior begins only when the corona of
the male squarely contacts the female. However, not all
head-on encounters result in mating; indeed, the probability of copulation in laboratory cultures generally
varies from 10 to 75% in Brachionus plicatilis, depending on the strain (Snell and Hawkinson, 1983).
This requirement for head-on contact by the male is
due to the presence of chemoreceptors in his coronal
region, which apparently respond only to a speciesspecific glycoprotein on the surface of the female (Snell
et al., 1995). Mating begins with the male swimming
circles around the female, skimming over the surface of
her lorica (Fig. 21). During this phase, the male maintains contact with the female with both his corona and
penis (this requires the male to remain in a slightly bent
position). After several seconds of circling, the male
attaches his penis to the female, usually in the region
of her corona, and loses coronal contact. After about
1.2 min of copulation in B. plicatilis (Snell and Hoff,
1987), sperm transfer is completed and copulation is
terminated when the male and female break apart and
swim away. Newborn B. plicatilis males only have
about 30 sperm and transfer two or three at each insemination (Snell and Childress, 1987).
3. Aging and Senescence
Rotifers have long been used as models of aging
and senescence. Jennings and Lynch (1928) conducted
FIGURE 21 Male mating behavior in brachionid rotifers. Arrows
indicate general swimming movements by male and female rotifers.
(Male not drawn to scale.)
the first major study of aging in rotifers using the monogonont Proales sordida. They found the mean lifespan
to be 8 days at 20°C, but a maternal effect was noted.
Offspring from older mothers had shorter lifespans and
more variability in their developmental rates and fecundities. Maternal effects were a theme expanded upon by
Lansing who, in a series of papers during the 1940s and
1950s, showed that parental age also influenced
longevity in the bdelloid Philodina citrina (see King,
1969 for a review). Lansing established “orthoclones”
by isolating the first offspring of parental females, then
the first offspring of F1 females, and so forth, yielding a
clone derived exclusively of the first-born females. Orthoclones derived from the offspring of older females
also were developed. This protocol is a powerful tool
for investigating effects on aging because the only difference among orthoclones is the age of their mothers.
In P. citrina, mean lifespans of 5-, 11-, and 16-day orthoclones were 23.8, 18.1, and 16.6 days, respectively.
Lansing’s general conclusion was that young orthoclones have increasingly longer lifespans, while older orthoclones have progressively shorter lifespans, resulting
eventually in clonal extinction. Rate of senescence,
Lansing hypothesized, was controlled by a maternal effect that was transmissible and cumulative. However,
the effect was reversible since the first-born females
from an old orthoclone outlived their parents. Based on
218
Robert Lee Wallace and Terry W. Snell
these and other studies, Lansing proposed that calcium
is the maternal factor responsible for what is now
known as the “Lansing Effect.” Rate of calcium accumulation and its importance in rotifer senescence became the reigning hypothesis of the day. Since Lansing’s
work, researchers have reported similar results in some
species, but not in others.
King (1983) re-examined Lansing’s data using life
table analysis and concluded that Lansing probably did
not observe an aging effect. King showed that shortlived lines derived from old parents reproduced earlier
and at higher rates in succeeding generations. Longlived lines derived from young parents delayed initial
reproduction to later age classes. King argued that the
Lansing effect actually resulted from shifts in fecundity
patterns that concentrated reproduction in a few age
classes in short-lived lines. Snell and King (1977)
demonstrated that concentrated reproduction in rotifers could shorten lifespan. Until more is known
about the physiological basis for the “Lansing Effect”,
King suggested that it be viewed with skepticism.
The pattern of reproduction also modifies
longevity (Snell and King, 1977). Female Asplanchna
brightwelli with high rates of reproduction concentrated early in life have shorter mean lifespans (2.5
days at 25°C) than females with slower reproduction
distributed over a greater portion of the life- span
(mean lifespan 4.5 days). The length of prereproductive and postreproductive periods were similar for
short- and long-lived individuals. However, length of
the reproductive period was about twice as long in
long-lived individuals. These data suggest that reproduction influences individual survival negatively, but
deleterious effects are minimized when the reproductive
period is spread over a greater portion of the lifespan.
Further, Rougier and Pourriot (1977) found that maternal age is negatively correlated with mictic female
production. When Brachionus calyciflorus females began reproducing at age 2 days at 15°C, 80 – 90% of
their daughters were mictic. By age 8 days, only 25%
of daughters were mictic, and reproduction by 12 day
old females yielded no mictic daughters. This decline in
mictic female production with age was linear in 2-, 6-,
and 10-day orthoclones.
Male lifespan has been examined far less thoroughly than that of females, but available data indicates that males live only about half as long as females
(Fig. 22) (King and Miracle, 1980; Snell, 1977).
Other experiments on aging in rotifers have indicated that vitamin E (a-tocopherol) can extend mean
lifespan about 10% at 23°C in the bdelloid Philodina
sp., but maximum lifespan is unaffected (Enesco and
Verdone-Smith, 1980). Vitamin E, at a concentration
of 20 g/mL, also was found to extend the lifespan
FIGURE 22
Female and male survival of three clones of
Brachionus plicatilis at 25°C. Clone LA is from LaJolla, CA; clone
MC is from McKay Bay near Tampa, FL; clone SP is from Castellon,
Spain. Note that the Y axis (survivorship %) is a log scale. (From
King and Miracle, 1980, with permission.)
in A. brightwelli (Sawada and Enesco, 1984). Of several other antioxidants tested, however, only thiazolidine-4-carboxylic acid (TCA) extended the mean life
span significantly, presumably by quenching free radical reactions.
Other factors that affect mean life span in A.
brightwelli are photoperiod (light:dark cycle, L:D), dietary restriction (Sawada and Enesco, 1984), and ultraviolet (UV) radiation (Enesco, 1993). Rotifers exposed
to a 6:18 or 0:24 L:D photoperiod showed an 18%
and a 22% increased life span, respectively, over a control photoperiod of 12:12 Dietary restriction has lifeextending effects in a wide variety of animals. In A.
brightwelli, two different modes of dietary restriction
yielded longer life spans. Reduction of food intake
increased life span by 14.2%, whereas intermittent
feeding produced an 11.8% longer life span. Ultraviolet irradiation in the 200 – 4800-J/m2 range shortened
life span significantly, and life span declined logarithmically as UV dose increased.
4. Population Dynamics
Studies of rotifer population dynamics attempt to
explain the causes of changes in population size. By
separating and quantifing the relative contributions of
reproduction, mortality, and dispersal, ecologists are
able to identify the factors regulating population size
and the determinants of average abundance. Because
some species are easily cultured and experimentally
manipulated, rotifers have proven to be useful models
for investigating the dynamics of animal populations.
Techniques also exist for investigating the dynamics of
natural rotifer populations, further broadening their
usefulness in ecological studies.
a. Life Tables The most rigorous approach to
studying population dynamics is lifetable analysis. Life
8. Phylum Rotifera
tables are age-specific records of all reproduction and
mortality occurring in a population. From these observations, vital population statistics can be calculated
such as intrinsic rate of increase (rm), net reproductive
rate (Ro), life span, generation time, reproductive value,
and stable age distribution. The general protocol for
gathering lifetable data on rotifers is to collect a cohort
of newborn females, isolate them in small volumes of
culture medium (ca. 1 mL), and make daily observations of their survival and reproduction. Maternal females usually are transferred daily to fresh medium
with a specific food level, and offspring are counted
and removed. A table is then constructed of age, number surviving in each age class, and number of offspring produced in each age class. At least 10 replicate
females are grouped into each population and probability of surviving to age x (lx) and age-specific fecundity (mx) are calculated.
The detailed observations required for life-table
analysis are usually only possible in laboratory populations and a number of genera have been examined
in that way, including several species of Brachionus
(Halbach, 1970; Pourriot and Rougier, 1975; King
and Miracle, 1980; Walz, 1987), Asplanchna (Gilbert,
1977; Snell and King, 1977), and bdelloids (Ricci,
1983). In certain cases, it is possible to perform life
table analyses on field populations (i.e., Floscularia
conifera, Edmondson, 1945). Although these and other
studies have examined survival and reproduction of
parthenogenetic females, survivorship curves for male
rotifers also are available for A. brightwelli (Snell,
1977) and B. plicatilis (King and Miracle, 1980).
The major environmental factors affecting age-specific survival and reproduction are temperature, food
quantity and quality, genetic strain, and reproductive
type (either amictic, or fertilized or unfertilized mictic
females). Increased temperature shortens survivorship.
In B. calyciflorus, for example, 50% of a cohort survive to age 16 days at 15°C, 11 days at 20°C, and 5
days at 25°C (Fig. 23). Age-specific fecundity also is
compressed at higher temperatures. At 15°C, fecundity
occurs at a low rate over 20 days. In contrast, at 25°C
reproduction occurs at a high rate and is compressed
into 10 days. Females produce a total of 13 offspring at
15°C, 16.6 at 20°C, and 12.9 at 25°C. The intrinsic
rates of increase (rm) calculated from these lx and mx
values are 0.34, 0.48, and 0.82 offspring per female
per day at the respective temperatures (Halbach, 1970).
The effect of food quantity on lx and mx schedules
is illustrated in Figure 24. Brachionus calyciflorus was
fed the green alga Chlorella at densities of 0.05 – 5
106 cells/mL at 20°C. The best survival occurred at
algal concentrations 0.5 106 and 1.0 106 cells/mL,
where mean life span was 9 days (Halbach and
219
Halbach-Keup, 1974). Life span decreased at lower
food levels, reaching 2.5 days at 0.05 106 cells/mL.
Lower life spans also were recorded at food concentrations above 1.0 106 cells/mL, probably the result of
accumulation of algal metabolic products that are
toxic to rotifers. Fecundity also peaked at 1.0 106
cells/mL, with a mean of 17 offspring per female. In
contrast, at 0.05 106 cells/mL, lifetime fecundity was
only 0.5 offspring per female whereas 3 offspring per
female were produced at 5.0 106 cells/mL.
Intraspecific differences in survival among strains is
illustrated in Figure 22. King and Miracle (1980) examined three strains of B. plicatilis and found that their
mean life spans at 25°C ranged from 6 to 13.5 days.
Because these strains were acclimated to, and tested in,
a common environment, the observed differences must
be genetic. Also shown in the same figure are male survivorship curves; interstrain differences are again apparent with male life spans.
The three different female types (amictic, and unfertilized and fertilized mictic) differ markedly in their agespecific survival and fecundity. In Brachionus urceolaris,
for example, amictic females live at 20°C an average of
9 days and produce 20 female offspring; unfertilized
mictic females live 9.5 days and produce 25 male offspring; and fertilized mictic females live 10 days and
FIGURE 23
Effect of temperature on age-specific survival and
fecundity of Brachionus calyciflorus. The left Y axis is the percent of
the cohort surviving (open circles) and the right Y axis is the offspring per female per hour (closed circles). (From Halbach, 1970,
with permission.)
220
Robert Lee Wallace and Terry W. Snell
FIGURE 25 The fecundity schedule and egg developmental times of
three types of Brachionus urceolaris females at 20°C with excess
food. Bottom panel: amictic females producing diploid female eggs;
middle panel: unfertilized mictic females producing haploid male
eggs; top panel: fertilized females producing diploid resting eggs
(solid circles). Arrows in the top panel represent the time of resting
egg attachment to fertilized mictic females(hrs). Each bar in the middle and lower panels represents one egg and the length of the bar
indicates the developmental time for that egg(days). (Modified from
Ruttner-Kolisko, 1974, with permission.)
FIGURE 24 Effect of food level on age-specific survival and fecundity of Brachionus calyciflorus. The left Y axis is the percent of the
cohort surviving (closed circles) and the right Y axis is the offspring
per female per hour (open circles). The food is Chlorella pyrenoidosa
provided at algal doses (AD) in the ( 0.05 – 5) 106 per ml/range.
(From Halbach and Halbach-Keup, 1974, with permission.)
produce four resting eggs (Fig. 25). The greatly reduced
fecundity of fertilized females is typical of other rotifer
species and is likely due to higher energy requirements
for resting egg formation (Gilbert, 1980a). Also reported in Figure 25 are development times for each egg
(time from egg extrustion to hatching) and the birth intervals between eggs. Birth intervals are shortest in the
middle of the reproductive period.
Individuals within a single rotifer population differing in their growth forms (e.g., spined and unspined)
can possess very different intrinsic rates of increase
under identical culture conditions. Stemberger (1988)
has shown that the rm value for spined and unspined
forms of Keratella testudo can differ significantly, but
the difference depends on food concentration (Fig. 26).
At low food levels, there is no significant difference in
rm for the two forms of this rotifer. At higher food concentrations, however, the unspined form had much
greater values of rm. Presumably, a similar phenomena
will be found in other species in which there are spined
and unspined forms (see Sections III.A.1, and III.C.4).
When life-history patterns of zooplankton are
compared, rotifers have higher rm values than either
cladocerans or copepods (Allan, 1976). The rm values
of these groups range from 0.2 to 1.6, 0.2 to 0.6, and
0.1 to 0.4 offspring per female per day, respectively.
Rotifers achieve their high population growth rates because of short development times, which more than
compensate for their small clutch sizes. Rotifers also
FIGURE 26
Intrinsic rates of population growth of spined (open
Figures) and unspined (closed Figures) Keratella testudo. Dashed
line, r 0. (Modified from Stemberger, 1988, with permission.)
8. Phylum Rotifera
show the greatest response to increased temperatures.
The high population-growth rates of rotifers may be an
adaptive response to predation, against which most
rotifer species have few defenses (Stemberger and
Gilbert, 1987a). As a result of these characteristics, rotifers are regarded as being able to quickly exploit new
conditions.
b. Dynamics of Field Populations Annual cycles of
natural rotifer populations have been characterized
for several species. One particularly thorough multiyear
study was conducted by Herzig (1987) in Neusiedlersee,
a shallow, well-mixed Austrian lake. Figure 27 shows
12 years of data for the abundance of 13 rotifer species.
Some species, like Keratella quadrata, have relatively
stable population sizes and occupy the lake nearly continuously. In contrast, abundances of other species, such
as Rhinoglena fertoensis, Filinia longiseta, and Hexarthra fennica, fluctuate much more. Fluctuation of
rotifer abundance over two or three orders of magnitude during a seasonal cycle is typical of many natural
populations.
Techniques have been developed to characterize the
dynamics of natural zooplankton populations (Edmondson, 1993). The instantaneous rate of increase of
a population (r) is the difference between instantaneous
birth (b) and death (d) rates
rbd
(1)
221
The value of r for a given population may be estimated from the equation
r
ln Nt ln N0
T
(2)
This procedure requires two successive estimates of
population size (No and Nt) separated by time interval
T. The estimate of r is based on several assumptions
about the population, including that it is growing exponentially with a stable age distribution. One can also
estimate the birth rate in this population by counting
the number of eggs carried per female. The finite hatching rate is
B
E
D
(3)
where E is the number of eggs per female and D
the developmental rate at a specific temperature. The
value E is determined directly from samples of the population and values of D by observing hatching of eggs
from a sample brought back to the laboratory. For example, Herzig (1983) reports values of D for several
species. When E and D have been calculated, the instantaneous birth rate, b, can be estimated from the
following equation by Palohiemo (1974)
b
ln (E 1)
D
FIGURE 27 Phenologies of several important rotifer species in Neusiedlersee, Austria. The X axis is the
season (winter, spring, summer, and autumn) and the Y-axis the rotifer abundance per cubic meter. (From
Herzig, 1987, with permission.)
(4)
222
Robert Lee Wallace and Terry W. Snell
With these data, it is possible to estimate the death
rate, d, by subtraction
dbr
(5)
The egg ratio technique makes it possible to predict
growth of natural rotifer populations where reproduction by amictic females is parthenogenetic. A few simple
measurements allow ecologists to estimate important
population parameters that summarize processes of
birth and death occurring in the population. This technique has proven useful for investigating the population
dynamics and secondary production of zooplankton
(e.g. Edmondson and Litt, 1987). However, before applying the egg ratio method, the works of Edmondson
(1993) and Palohiemo (1974) should be consulted.
The dynamics of natural rotifer populations are affected by a number of environmental factors including
temperature, food quantity and quality, exploitative and
interference competition, predation, and parasitism.
Temperature is a major factor affecting fertility, mortality, and developmental rates. In general, higher temperatures do not increase the number of offspring produced per female, but instead, shorten birth intervals by
decreasing developmental time (Edmondson, 1993;
Pourriot, 1965). Life span also is reduced, but the net
effect of higher temperatures is an increased population
growth rate. In laboratory studies, higher temperatures
and elevated food levels combine synergistically to increase significantly mean rate of population increase (r)
of Brachionus calyciflorus (Fig. 28). Other abiotic environmental factors, such as oxygen concentration, light
intensity, and pH, also influence rotifer population dynamics (see Hofmann, 1977 for a review).
Food availability is a major biotic factor regulating
rotifer population growth. Species have markedly different food requirements for reproduction. Threshold food
concentration, the food level where population growth
is zero, was determined for eight species of planktonic
FIGURE 28
Synergistic effects of temperature and food density on
mean rate of population increase (rm) of Brachionus calyciflorus.
(From Starkweather, 1987, with permission.)
FIGURE 29
Threshold food levels where population growth rate
(rm) 0. Food concentration is algal dry mass/mL; rm is offspring per
female per day. The solid line indicates food concentrations required
to maintain a population growth of zero. Values above the dashed
line (positive rm) indicates increasing population densities; values
below the solid line (negative rm) indicate decreasing population
densities. Species code: Kc Keratella cochlearis, Ke Keratella
earlinae; Kcr, Keratella crassa; So, Synchaeta oblonga; Sp, Synchaeta
pectinata; Bc, Brachionus calyciflorus; Pr, Polyarthra remata; and Ap,
Asplanchna priodonta. (From Stemberger and Gilbert, 1985b, with
permission.)
rotifers by Stemberger and Gilbert (1985). Small species
like Keratella cochlearis had lower threshold food concentrations than larger species like Asplanchna priodonta or Synchaeta pectinata (Fig. 29). The logarithm
of the threshold concentration was positively related to
the logarithm of rotifer body mass, so the smallest
species had the lowest food thresholds. The food concentration required to support 50% of the maximum
population growth rate (rmax/2) varied 35-fold among
the eight rotifer species and also was positively related
to body mass. Small rotifer species, therefore, appear
best adapted to food-poor environments because those
species possess low food thresholds. Larger species, in
contrast, are better adapted to food-rich environments,
where they have higher reproductive potentials.
c. Genetic Variation Our knowledge of rotifer genetics is in its infancy. An early worker, Hertel (1942) observed strong inbreeding depression in five strains of
Epiphanes senta after one generation of inbreeding
by self-fertilization. Net fecundity (R0) was reduced
70 – 80% in the F1 generation and continued to decline
slowly in F2 and F3 generations. Birky (1967) found similar results in Asplanchna brightwelli, with a 25 – 45%
reduction in percentage of resting eggs hatching after one
generation of selfing. Inbreeding increases homozygosity,
thus potentially revealing deleterious recessive alleles previously masked by dominance. Inbreeding depression
suggests that substantial amounts of genetic variability
exist in natural rotifer populations.
Electrophoretic analysis of allozymes may be used
to characterize genetic variation in natural populations.
8. Phylum Rotifera
Although application of these techniques to rotifers has
been very limited, allozymes have been used to distinguish strains. Snell and Winkler (1984) analyzed five enzymes representing seven loci in 17 strains of Brachionus
plicatilis from all over the world. This work showed no
relationship between geographic proximity and genetic
similarity. In the same species, Serra and Miracle (1985)
found low allozyme variation, with only one or two
electrophoretic patterns present in each population.
These researchers suggested that this low variability resulted from intense inbreeding and interclonal competition, both of which lead to stabilization of just a few
highly adapted genotypes. Although genetic variance
within populations was low, Serra and Miracle found
substantial variance between populations. In contrast,
King and Zhao (1987) demonstrated that several electromorphs were present in each collection of B. plicatilis
from Soda Lake in Nevada, rather than a few dominant
types as reported by Serra and Miracle (1985). King and
Zhao also observed no seasonal succession of genotypes
and concluded that B. plicatilis in Soda Lake follow a
pattern of incomplete genetic discontinuity. In this pattern, several genotypes persist throughout the year, but
their frequencies increase or decrease in response to environmental changes (King, 1972). Allozyme variation
was analyzed in five populations of the bdelloid Macrotrachela quadricornifera using six enzymes (Pagani,
et al., 1991). Relatively high polymorphism was found,
with only 27% of alleles shared by all strains.
A variety of perspectives on the genetic structure of
rotifer populations have been offered. Ruttner-Kolisko
(1963) argued that most rotifer populations are highly
homogeneous due to founder effects and strong selection
eliminating all but a few genotypes. As a result, bisexual
reproduction in rotifers is characterized by extreme inbreeding. Such a genetic structure produces fragmented
gene pools where geographically separate populations
also are isolated genetically. Birky (1967) disagreed with
this view, stating that the substantial heterozygosity revealed by inbreeding experiments suggests that rotifer
populations are not genetically homogeneous. The great
morphological similarity among geographically separated populations (Pejler, 1977a; Dumont, 1983) further
supports the idea that rotifer gene pools are better integrated than Ruttner-Kolisko proposed.
Seasonal changes in rotifer population structure
have been characterized by King (1972) who proposed
three models; one based on physiological adaptation;
and two based on genetic changes. Working with Asplanchna girodi in a small pond in Florida, King (1977,
1980) concluded that the complete genetic discontinuity
model best described his observations. Asplanchna
girodi actually had a succession of genetically distinct
populations, each hatching from resting eggs, complet-
223
ing their life cycle, and returning to dormancy as the
next population was emerging. Little overlap occurred
between populations, thus making them completely discontinuous. Competition (Snell, 1979) and the presence
of cyanobacteria (Snell, 1980) were major ecological
factors promoting succession among these rotifer populations. The population structure differs from the incomplete genetic discontinuity structure that King and
Zhao (1987) reported for B. plicatilis in Soda Lake.
Experimental data from interpopulation crosses
also illuminate the genetic structure of rotifer populations. Some strains of rotifers can mate successfully
with strains from widely separated habitats, while others such as Asplanchna brightwelli (Birky, 1967; King,
1977) are reproductively isolated.
Behavioral reproductive isolating mechanisms are
well developed in rotifers and can be used to define
species boundaries (Snell, 1989). Gilbert (1963b) investigated this phenomenon in Brachionus calyciflorus
and found that mating behavior was strictly speciesspecific; B. calyciflorus males failed to mate with females of B. angularis, B. quadridentatus, Synchaeta or
Euchlanis. Although Snell and Hawkinson (1983) reported no behavioral reproductive isolation among
temporally separated populations of B. plicatilis from
the same bay, reduced probabilities of copulation
among spatially and geographically isolated populations revealed incipient behavioral isolation.
Reproductive isolating mechanisms are not always
effective barriers to gene flow. Pejler (1956) argued that
the morphological intermediates of Polyarthra vulgaris
and P. dolichoptera he found in several Swedish tarns
were the result of interspecific hybridization. RuttnerKolisko (1969) provided some direct evidence of interspecific hybridization when she successfully hybridized
B. urceolaris and B. quadridentatus. Resting eggs were
formed from this cross (in both directions), but they
could be hatched only when female B. urceolaris were
crossed with male B. quadridentatus. Asymmetry in hybridization is typical of closely related species and suggests that reproductive isolation between B. urceolaris
and B. quadridentatus has yet to be completed.
C. Ecological Interactions
1. Foraging Behavior
Although the diets of some rotifers are highly specialized (Bogdan and Gilbert, 1987), many species consume a wide variety of both plant and animal prey and
may be described as generalist suspension feeders
(Walz, 1997). For example, Asplanchna species generally are considered predatory, yet their main food
often can be large algae. In contrast, the “herbivorous”
Brachionus and Ptygura will also eat small ciliates.
224
Robert Lee Wallace and Terry W. Snell
Thus, distinctions between predator and herbivore in
rotifers often are not clear; it is more constructive to
consider the ways in which rotifers encounter and
process food items rather than what they eat.
Research on food selection by rotifers began by
correlating the density of natural populations of rotifers to the food available (Edmondson, 1965) and by
determining food selectivities using laboratory cultures
and/or natural populations (Bogdan and Gilbert, 1982,
1987; Salt, 1987). In the latter studies, natural foods
(i.e., algae, bacteria, yeast) or artificial materials (e.g.,
latex microspheres) are used to observe rotifer feeding
behavior and to calculate feeding rates. Natural foods
have been labeled using radioisotopes, but loss of label
from the food can result in significant errors. However,
even the simple procedure of adding a few drops of
food suspension to a Petri dish with rotifers may lead
to interesting observations concerning how rotifers
process food. For example, using powdered carmine,
Wallace (1987) demonstrated that adult Sinantherina
socialis do not act independently of one another when
in a colony. Instead, the coronae of the animals in a
large section of the colony all face in the same direction
for several minutes. Animals exhibiting this behavior
are called an array. Arrays determine where the dominant water currents flow to the colony, how the water
currents flow around the colony, and where the currents leave the colony.
Rotifers feed in ways that are directly related to
their general life history. Although most planktonic rotifers, such as Asplanchna, Brachionus, Polyarthra, and
Rhinoglena, swim at similar rates through comparable
areas of the water column, the ways in which they
encounter and capture food can be quite different. Brachionus uses its swimming currents to sweep small
food particles into the buccal region for processing; this
is often termed filter or suspension feeding.
Asplanchna, on the other hand, does not use its swimming currents to gather food. Once this rotifer contacts
a potential food item with its corona, it may or may
not attempt to ingest the item based on such factors as
hunger level and the size and type of prey (Gilbert and
Stemberger, 1985b; Salt et al., 1978). Some benthic
rotifers creep along algal filaments and feed by piercing
the filament and sucking the cytoplasm from the cells;
examples are Notommata copeus (Clément et al.,
1983) and Trichocerca rattus (Clément, 1987). Sessile
rotifers capture food in one of two ways, depending
on the family. Members of the family Flosculariidae
(e.g., Floscularia, Ptygura, Sinantherina) create feeding
currents in a manner similar to the planktonic suspension feeders. In contrast, all collothecid rotifers are ambush raptors (e.g., Collotheca, Stephanoceros). Once a
prey enters the infundibular region of the corona of
these rotifers, long setae (Collotheca) or arms (Stephanoceros) fold over the prey, capturing it much like the
action of a Venus flytrap. All members of the family
Atrochidae lack setae (e.g., Cupelopagis). In these
forms, an enlarged, sometimes sheetlike corona folds
over prey to capture them. In both cases, prey are
pushed through the rotifer’s mouth and into the
proventriculus where they are stored until the mastax
transfers them into the stomach. When prey are particularly abundant, several live prey may be seen in the
proventriculus.
Once food is captured, the process of ingestion begins, but not all captured food items are consumed. By
observing individual Brachionus calyciflorus in various
densities of suspended food particles, Gilbert and
Starkweather (1977, 1978) showed that this rotifer regulated its ingestion of food particles by three different
mechanisms. First, rotifers used pseudotrochal cirri to
screen certain large particles away from the mouth.
Second, particles collected by the corona may be rejected later by cilia within the buccal field. Finally, particles that gain entrance to the oral cavity may be returned to the buccal field by action of oral cilia for
subsequent rejection. The mastax also may actively reject particles. Microphagus rotifers, such as Brachionus
and Ptygura, use the mastax to push food (usually
larger particles) out of the oral cavity. Members of the
genera Asplanchna and Asplanchnopus also use their
trophi to remove materials from their stomachs, but
the materials removed by these raptors are empty carapaces of hard-bodied prey such as cladocerans. Occasionally, predatory rotifers such as Cupelopagis, may
become so engorged that attempting to ingest another
prey item results in the loss, from the proventriculus, of
one previously captured.
2. Functional Role in the Ecosystem
Freshwater zooplankton are dominated by three
taxonomic groups (protozoans, rotifers, and microcrustaceans), yet it is the microcrustaceans that usually receive the most attention from researchers. Such a disparity is probably to be expected for two reasons, one
significant and one trivial. First, microcrustaceans commonly account for a greater proportion of the total
zooplankton biomass than either protozoans or rotifers. Second, microcrustaceans are easier organisms to
manipulate and observe in laboratory situations. However, although rotifers usually have a smaller standing
biomass than microcrustaceans, their high turnover
rate makes them very important to the trophic dynamics of freshwater planktonic communities (e.g., Bogdan
and Gilbert, 1982, Neil, 1984).
Feeding rates of zooplankton generally are referred
to as filtration or clearance rates and are measured as
8. Phylum Rotifera
microliters of water cleared of a certain food type per
animal per unit time (i.e., L/animal/h). Usually rotifer
clearance rates are lower than those of cladocerans
and copepods, although the rates depend heavily on
food type, temperature, and animal size (Bogdan and
Gilbert, 1982). For most rotifers, clearance rates are
commonly between 1 and 10 L/animal/h, whether determined in the laboratory or in the field. However, a
few species can achieve levels exceeding 50 L/animal/h (e.g., Bogdan and Gilbert, 1987).
Using estimates of clearance rates, Starkweather
(1987) noted that even moderately sized rotifers with
body volumes of about 10 3 L process enormous
amounts of water with respect to their size: 103 times
their own body volume each hour! Ingestion rates (biomass consumed per animal per unit time) also are very
high for rotifers. An adult rotifer may consume food
resources equal to 10 times its own dry weight per day.
Therefore, by having assimilation efficiences (i.e., assimilation divided by ingestion) of between 20 and
80%, rotifers convert a good deal of their food to animal biomass, which may be passed on to the next
trophic level (Starkweather, 1980, 1987).
Although microcrustaceans generally have higher
clearance rates than rotifers (about 10 – 150 L/animal/h for cladocerans and 100 – 800 L/animal/h for
copepods) rotifers can exert greater grazing pressure on
phytoplankton than some small cladocerans. Bogdan
and Gilbert (1982) report that in a small eutrophic lake,
Keratella cochlearis accounted for about 80% of the
community grazing pressure on small algae during the
year. These two workers also showed that K. cochlearis
had filtering rates about 5 – 13 times higher than that
for the cladoceran Bosmina longirostris (Gilbert and
Bogdan, 1984). Therefore, under certain conditions, rotifers may be important competitors to small, filter-feeding microcrustaceans and are important in nutrient
recycling in aquatic systems (Makarewicz and Likens,
1979). Furthermore, rotifers can alter the species composition of algae in certain systems. Schlüter et al.
(1987) showed that intense feeding by Brachionus
rubens can cause a shift in the dominant algal species
from Scenedesmus to a spined algae, Micractinium. Apparently, this shift is based on the inability of B. rubens
to consume algae with protective spines.
Although generally not ingested, cyanobacteria are
increasingly recognized as playing an important role in
determining zooplankton species composition. Large
cladocera are more sensitive to the cyanobacteria Anabena affinis than rotifers (Gilbert, 1990). The mechanism for this differential sensitivity to cyanobacteria
toxicity is based on different tendencies to ingest
cyanobacterial filaments and different physiological
tolerances of the toxin (Kirk and Gilbert, 1992).
225
Having the ability to reproduce rapidly, rotifers
may account for 50% or more of the zooplankton production, depending of the prevailing conditions (Herzig,
1987). This production, in turn, can be an important
food source for other rotifers, Asplanchna (Gilbert and
Stemberger, 1985b), cyclopoid and calanoid copepods
(Williamson, 1983), malacostracans (Mysis, Threlkeld
et al., 1980), insect larvae (Chaoborus, Moore, 1988;
Moore and Gilbert, 1987), and fish (Hewitt and
George, 1987).
Abundance and species composition of rotifers often reflect the trophic status of lakes. For example,
Hillbricht-Ilkowska (1983), Walz et al. (1987), and
others have reported changes in the maximal totalpopulation density of several orders of magnitude
when lakes were subject to intense eutrophication. Individual species sometimes undergo dramatic population changes during those periods. Walz et al. (1987)
showed that the density of Asplanchna in Lake Constance increased its maximum population levels 280fold over a period of 28 years. Population declines also
have been seen. Edmondson and Litt (1982) indicated
that Keratella cochlearis was abundant during the years
when Lake Washington had elevated concentrations of
dissolved phosphorus, low water transparency, and
high algal densities. However, as these waterquality
parameters improved, the population of this rotifer
declined dramatically. Overall, there was at least a
20-fold increase, followed by a decline during a period
of 15 years.
Studies of the interactions of rotifers with other
organisms will probably continue to receive attention
in the future, especially predator – prey interactions
(Gilbert, 1966 1967; Salt, 1987; Williamson, 1983), interference competition between other herbivorous zooplankton and rotifers (Burns and Gilbert, 1986a,
1986b; Gilbert and Stemberger, 1985a), the toxic effects of cyanobacteria (Snell, 1980; Starkweather and
Kellar, 1987), mate recognition (Snell et al., 1995), and
life-history strategies (Walz, 1995). One underutilized
tool for the study of energetics and trophic interactions
is the chemostat, which, unlike batch cultures, maintains constant experimental conditions (Boraas and
Bennet, 1988; Walz, 1993).
3. Competition with Other Zooplankton
Rotifers, cladocerans, and copepods often compete
for limited food resources and, in general, rotifers are
relatively poor exploitative competitors because their
clearance rates are usually many times lower than those
of daphnids (see Section III.C.2). Rotifers also have a
more limited size range of particles they can ingest
compared to cladocerans and are less resistant to starvation (Kirk, 1997). Most rotifers eat algal cells in the
226
Robert Lee Wallace and Terry W. Snell
4 – 17 m range and are much less efficient at ingesting
smaller or larger cells (Gilbert, 1985b). In contrast,
most cladocerans eat algal and bacterial cells in the
1 – 17 m range, and some can ingest much larger cells
(Hall et al., 1976; Bogdan and Gilbert, 1982). Thus,
cladocerans generally have broader food niches than
rotifers in terms of food type and size, and through direct competition may suppress rotifer population
growth. This outcome may be reversed, however, when
a sufficient quantity of suspended sediments is present
in a lake (Kirk, 1991).
Exploitative competition between rotifers and
daphnids is readily demonstrated when these zooplankton are grown in single and mixed cultures. Brachionus
calyciflorus and Daphnia pulex both grow well on the
alga Nannochloris oculata in single species cultures.
When both species are present, however, Daphnia removes an increasingly larger proportion of algal cells
until the rotifers gradually starve to extinction after
2 – 3 weeks (Fig. 30). Daphnia is unaffected by the
presence of rotifers.
Population growth of certain rotifers also is inhibited by Daphnia through interference competition
(Burns and Gilbert, 1986a, b; Gilbert and Stemberger,
1985b; Gilbert, 1989a). For example, in the presence of
any one of four Daphnia species, Keratella cochlearis
suffers mechanical damage (i.e., killed, wounded, or
loss of eggs) when swept into the branchial chamber of
the daphnids. No species differences were detected
among the Daphnia used, but the rate at which rotifers
were killed was directly proportional to daphnid body
length. Keratella also was found in the guts of some
Daphnia, documenting a new pathway for trophic interactions. Daphnia have a major impact on Keratella
populations when the cladocerans are larger than 2 mm
and present in densities of 1 – 5 individuals/L. Similar
effects have been reported for short-term bottle experiments involving the cladoceran Scapholeberis kingi and
the rotifer Synchaeta oblongata (Gilbert, 1989b). Of
course, interference and exploitative competition may
occur simultaneously (Gilbert 1985b, 1988).
Predation is another important regulatory factor in
rotifer population dynamics, as rotifers are prey for
several aquatic predators including protozoans, other
rotifers, insect larvae, cladocerans, copepods, and
planktivorous fish. From many studies we know that
predation affects rotifer population dynamics both directly (by contributing to mortality) and indirectly as a
selective force shaping rotifer morphology, physiology,
and behavior (see the next section). However, one area
that deserves more study is the predatory interactions
of protozoans and rotifers (e.g., protozoans as
predators — Finlay et al., 1987 and Nogrady et al.,
1993; rotifers as predators — Arndt, 1993; and Gilbert
and Jack, 1993).
4. Predator – Prey Interactions
FIGURE 30
Competition between Brachionus calyciflorus and
Daphnia pulex. Brachionus and Daphnia were grown in single
species (closed symbols) and mixed-species (open symbols) batch
cultures at 20°C, daily renewed with 5 106 Nannochloris cells/mL.
Population size (Y axis) is the number of individuals in the 80 mL
culture. Error bars ( 1 se) are visible when they exceed the size of
the symbols. (From Gilbert, 1985, with permission.)
Rotifers possess several types of defensive mechanisms that have apparently evolved in response to
predation. These mechanisms include morphological
features, escape movements, and other ways of avoiding potential predators (see Stemberger and Gilbert,
1987a; Williamson, 1983, 1987).
Most rotifers are transparent and quite small,
some comparable in size to protozoans (ca. 60 – 250
m long). While these features benefit limnoplanktonic
rotifers by reducing their visibilty to fish, a small body
size renders rotifers more vulnerable to invertebrate
predators, which feed by touch rather than by sight.
Many rotifers produce a thickened integument (lorica)
and/or spines and other projections, or carry their
eggs, all of which have been shown to reduce the ability of predatory zooplankton to prey upon them (e.g.,
Gilbert, 1966, 1967, 1980b; Stemberger and Gilbert,
1984b). Spines are produced in some rotifers (e.g.,
Brachionus calyciflorus, Figs. 12 and 31; Keratella
cochlearis; Keratella testudo, Fig. 26) in response to a
build-up of soluble substances released by invertebrate
predators such as Asplanchna and copepods (Gilbert,
1967; Stemberger and Gilbert, 1984b; Stemberger,
8. Phylum Rotifera
1984). We now know that exposure to a water-soluble
factor secreted by the predatory rotifer Asplanchna or
the copepods Epischura, Mesocyclops, and Tropocyclops induces de novo spine formation or spine lengthening in several planktonic rotifers (Stemberger and
Gilbert, 1987a, 1987b) (see Section III.A.1). Rotifers
for which such polymorphisms have been observed include Brachionus calyciforus, B. bidentata, B. urceolaris, Filinia longiseta passa, Keratella cochlearis, K.
slacki, and K. testudo. The adaptive significance of
spined morphotypes is a significant reduction in capture and ingestion by invertebrate predators by making the rotifer more difficult to manipulate and swallow. The presence of spines on B. calyciflorus is a good
example of the phenomenon (Fig. 12). The way this
works is as follows. When disturbed by a potential
predator, B. calyciflorus will retract its corona and by
doing so increases the hydrostatic pressure within its
pseudocoelom. The elevated pressure causes the posterolateral spines to swing forward and outward
(Fig. 31). Thus, the apparent body volume is larger.
For Asplanchna, this increase in the size of its prey is
sufficient to prevent ingestion after capture. After a period of time during which Asplanchna attempts to
swallow B. calyciflorus (usually 60 sec), the predator
will release (reject) the prey which then swims away
unharmed. Thus, the invertebrate predator releases a
biochemical cue (an allelochemic) that initiates a developmental change in subsequent generations of the
rotifer (spine production) reducing the effect of predation. Some forms of K. cochlearis also possess posterior spines, which make them four times more likely to
be rejected after capture by A. girodi than unspined
forms (Stemberger and Gilbert, 1984b). Most rejected
prey swim away unharmed. However, occasionally
spined forms are swallowed by Asplanchna or a similar genus (Asplanchnopus) and the spines become
lodged in the pharynx or esophagus of the predator.
FIGURE 31 Extension of the postero-lateral spines in Brachionus
calyciflorus. (Modified from Koste, 1976, with permission.)
227
FIGURE 32 Filinia (lateral view). This rotifer possesses long spines
which are used in making rapid jumps to escape predators. (Modified
from several sources.)
Presumably, both predator and prey die when that
happens.
Spine production has additional consequences beyond predatory avoidance. Stemberger (1988) has
demonstrated that, in the absence of predators, reproductive effort (survivorship and fecundity) and hydrodynamic characteristics (sinking rates and swimming
speeds) vary in spined and unspined Keratella testudo.
These relationships were further complicated by food
concentration (see Section III.B.4.a, and Fig. 26).
Transparent mucus sheaths produced by some
planktonic rotifers (e.g., Conochilus and Lacinularia
species) appear to function in a similar way to spines.
The sheaths make the rotifers effectively larger and,
therefore, less vulnerable to invertebrate predators
(e.g., Asplanchna and predatory copepods) without
making them more visible to fish (Gilbert, 1980b;
Stemberger and Gilbert, 1987a, 1987b; Wallace, 1987).
Three rotifer genera can often escape predators by
making rapid jumps using a variety of appendages: long
spines (Filinia, Fig. 32), setous armlike appendages
(Hexarthra, Fig. 13), and paddles (Polyarthra, Fig. 33)
(see Section II.C.1). Many rotifers assume a passive
posture, displaying what has become known as the
“dead-man response,” rather than fleeing when predators attack (e.g., Asplanchna, Brachionus, Keratella,
Sinantherina, and Synchaeta). This simple behavior is
merely a retraction of the corona into the body and
passive sinking. Contraction of the corona stops the animal from swimming, thus eliminating the vibrations it
produces in the water that may be detected by the
predator. In addition, this behavior may make the rotifer more difficult to grasp in its turgid state. In Sinantherina spinosa, this passive posture exposes a group of
228
Robert Lee Wallace and Terry W. Snell
FIGURE 33
Polyarthra. This rotifer possesses long paddlelike appendages which are used in making rapid jumps to escape predators.
(Modified from several sources.)
cases, it appears that parasites can cause the demise of
an entire population within a few days (Edmondson,
1965).
The sporozoan parasite, Microsporidium (Plistophora aerospora) frequently infects planktonic rotifers
possessing thin loricas (Ruttner-Kolisko, 1977a). For
example, members of the genera Asplanchna, Brachionus, Conochilus, Epiphanes, Polyarthra, and Synchaeta have been reported to be parasitized. Apparently, water temperature is an important mediating
factor in the spread of the parasite as infection rate
drops off at water temperatures below 20°C. In infected rotifers, the pseudocoel of the animal becomes
nearly filled with sausage-shaped cysts of Microsporidium (Fig. 34).
Several workers have described endoparasitic fungi
which attack soil rotifers of the genera Adineta and
Philodina. These reports describe three avenues of parasitic attack: adhesion, ingestion, and injection. Tzean
and Barron (1983) have described one fungus that forms
peglike, adhesive appendages on both small conidia
(spores, ca. 30 m long) and long vegetative hyphae.
small spines on its ventral body surface, which may
function in defense against planktivorous fish (see Wallace, 1987).
To date, only one species of rotifers, Sinantherina
socialis, has been shown as unpalatable to small zooplanktivorous fishes (Felix et al., 1995). While neither
the nature nor the location of the unpalatability factor(s) is known, these authors speculate that this colonial species possesses an unidentified chemical held in
the glandlike “warts” which are located at the anterior
end of the animals.
Defenses against predators like spines, mucus
sheaths, thickened loricas, and escape movements are
energetically demanding. Some species, like Synchaeta
pectinata, are not well defended against predators, but
have evolved very high, maximal population growth
rates that offset mortality from predation. Avoiding potential predators in space and/or in time is another simple yet effective defense mechanism against predators.
Some rotifers occupy the habitat at a different time of
year than an important predator, migrate vertically or
horizontally in the habitat, or live in zones with low
oxygen concentration, thereby missing predatory pressures altogether.
5. Parasitism on Rotifers
The importance of parasites in controlling population density in rotifers has not been examined thoroughly, although a few studies have correlated parasitic
infection with a decrease in population density of
planktonic species (Ruttner-Kolisko, 1977a). In certain
FIGURE 34
Photomicrograph of Brachionus infected
Microsporidium within its body cavity (pseudocoel).
with
8. Phylum Rotifera
FIGURE 35
Fungal parasites of rotifers. (A) Four rotifers trapped
by adhesive pegs on the vegetative hyphae of Cephaliophora; (B) unicellular, assimilative hyphae of Triaculus; and (C) zoosporangium of
Haptoglossa with two evacuation tubes and ten encysted zoospores.
(Bars ⬇ 100 m.)
Once the adhesive pegs attach to a rotifer, they germinate and rapidly colonize the pseudocoel (Fig. 35A). At
least three different genera of fungi produce spores
(Fig. 35B) that initiate parasitic attack when ingested
(e.g., Barron, 1980a; Barron and Tzean, 1981). A third
avenue of infection occurs via hypodermic injection of a
vegetative cell into the rotifer (Barron, 1980b; Robb and
Barron, 1982). Once inside the host, these fungal cells
grow into assimilative hyphae, producing more infective
cells either inside or outside the rotifer (Fig. 35C).
6. Aquaculture
In freshwater communitites, rotifers serve as food
for several invertebrate predators which, in turn, are
consumed by many economically important fish. They
also are used directly as food by many planktivorous
adult fish (O’Brien, 1987). Perhaps the most important
ecological link between rotifers and fish is the consumption of rotifers by larval fish. At an early age,
229
most fish larvae feed on microzooplankton, rotifers being one of the most important components of the diet.
In general, rotifers are highly nutritious and their biochemical composition can be further improved by specialized diets (Watanabe et al., 1983). Many fish are
well adapted to locate, pursue, capture, and ingest rotifers, and rotifers are easy prey because they swim
slowly and frequently lack predator defenses (Stemberger and Gilbert, 1987a).
Aquaculturists have exploited this important relationship between planktivorous fish and their rotifer
prey in intensive aquaculture systems. This field has developed into a major technical discipline in several
countries, most notably Israel, Japan, and China. Most
of this work has the practical goal of determining the
correct biotic and abiotic factors necessary to maintain
mass cultures of rotifers (usually Brachionus plicatilis)
as food organisms for larval shrimp and fish (Lubzens,
1987; Lubzens et al., 1989).
Most systems for mass culturing of rotifers are
simple batch cultures capable of producing kilogram
quantities of rotifer biomass each day (Hagiwara et al.,
1997). A substantial amount of effort has been devoted
to identifying optimal culture conditions for the most
commonly grown species (Brachionus rubens and B.
calyciflorus in freshwater and B. plicatilis in saltwater).
For mass cultures of B. rubens raised at 20°C, the optimal pH is 6.0 – 8.0, with upper and lower lethal limits
of 9.5 and 4.5, respectively (Schlüter, 1980). Oxygen
levels as low as 1.15 mg/L sustained good growth, but
0.72 mg oxygen/L was strongly limiting (Schlüter and
Groeneweg, 1981). Schlüter (1980) also tested eight algal species and found that Scenedesmus costato-granulatus, Kirchneriella contorta, and Chlorella fusca produced the best rotifer growth. Algal concentrations of
70 mg/L dry weight of Scenedesmus were optimal. Brachionus rubens reproduction in mass cultures is inhibited by ammonia levels of 3 – 5 mg NH3-N/L (Schlüter
and Groeneweg, 1985). Because about 50% of the nitrogenous wastes excreted by Brachionus is ammonia,
waste product removal is a serious consideration in
mass culture design (Hirata and Nagata, 1982).
Although rotifers have been important in marine
fish aquaculture for many years (Lubzens, 1987), freshwater rotifers are just beginning to be used. Nauplii of
brine shrimp (Artemia) have been used extensively as
larval feed, but these marine organisms do not live more
than about 6 h in freshwater, and their decomposition
contributes to the organic load in culture tanks. A freshwater organism is preferred for freshwater aquaculture,
and rotifers fill this role well. Rotifers also are being
used as food for young fish larvae too small to begin
feeding on Artemia. For these reasons the use of rotifers
in freshwater aquaculture is expected to grow.
230
Robert Lee Wallace and Terry W. Snell
D. Evolutionary Relationships
The phylogenetic position of the eight tradition
“pseudocoelomate” phyla represents a problem that
continues to generate debate among workers. Within
this larger question several opinions have been expressed concerning the evolution of rotifers, including
some that propose a polyphyletic origin of the phylum.
One theory suggests that rotifers evolved from an ancestor possessing a fluid-filled body cavity (Lorenzen,
1985), perhaps a regressed coelom, while another proposes that rotifers were derived from an ancestral acoel
turbellarian (phylum Platyhelminthes, Chapter 6) (Hyman, 1951). We believe that the evidence favors the
FIGURE 36
Phylogenetic relationships within phylum Rotifera at
the class level. Illustrated here are the three different hypotheses
regarding the relationship of acanthocephalans and rotifers. (A) The
traditional hypothesis based on an analysis of morphological data
concludes that acanthocephalans are a sister group to rotifers
(Melone et al., 1998). (B) A recent hypothesis based on an analysis of
molecular data (nuclear 18S rRNA and mitochondrial 16 S rRNA)
concludes that acanthocephalans are a highly modified sister group
to bdelloid rotifers (Garey et al., 1998). (C) Results of recent research
of molecular data (nuclear gene: 82 kd, heat-shock protein)
concludes at acanthocephalans are highly modified rotifers related to
the sister group of bdelloids plus monogononts (Mark Welch, 2000).
Numbers refer to changes in character state as gains or losses. Character states are numbers as follows: (1) intracytoplasmic lamina; (2)
females with uterine bell, ciliated epidermal cells absent, cephalic
gland absent; (3) ciliated corona present, mastax with trophi present,
retrocerebral apparatus present and paired, dorsal antenna present;
(4) Eurotatoria: urinary collection bladder present, manubrium
present in trophi, paired retrocerebral apparatus fused into a single
unit; (5) male genital ducts U-shaped, penis absent; (6) male haploidy, single gonads; (7) adult capable of a type of diapause, loss of
male, stomach syncytial, fulcrum absent in trophi; and (8) specialized
parasitic features.
view that rotifers and platyhelminthes are related; these
phyla share several common features including protonephridia, a ciliated integument which possesses mucus glands, and a cerebral eyespot with a pigment cup
(Wallace et al., 1996; Garey et al., 1998).
The most critical issue regarding the origin of the
Rotifera is the question of its relationship to phylum
Acanthocephala. Presence of the intracytoplasmic lamina within a syncytial epidermis as well as other morphological features in both phyla suggests that they are
closely related even though acanthocephalans are
5 – 1500 times larger in size than the largest rotifer
(Wallace et al., 1996). Lorenzen (1985) argues that
structures identical to the rostrum and lemnisci of
acanthocephalans are found in bdelloids, thus making
the two monophyletic. However, Ricci’s (1998a) recent
critique argues against homology of these structures
and a recent cladistic analysis of some 60 morphological characters rejected Lorenzen’s thesis that acanthocephalans and bdelloid rotifers are related (Melone et
al., 1998b). Nevertheless, molecular analyses of 18S
rRNA genes from a few well-known species place the
acanthocephalans firmly within the Rotifera. One view
places the acanthocephalans as a highly modified sister
group to the bdelloids (Garey et al., 1996, 1998), while
another places them as highly modified rotifers related
to the sister group of bdelloids and monogononts
(Mark Welch, unpublished observations). If additional
genetic analysis confirms these studies, the definition of
what constitutes the phylum Rotifera will need to be
completely re-interpreted. A composite overview of
these hypotheses is illustrated in Figure 36.
IV. COLLECTING, REARING, AND PREPARATION
FOR IDENTIFICATION
A. Collections
Collecting rotifers does not require complex or expensive equipment. One can almost always collect several species of planktonic rotifers by towing a finemesh (25 – 50 m) net through any body of water.
[N.B.: Nets with greater mesh sizes tend to miss smallbodied forms.] Productive lakes and ponds usually provide especially good sampling sites. More elaborate
equipment such as closing nets and Clarke – Bumpus
samplers work well, but are not necessary unless required by a specific experimental protocol. Water collected by discrete sampling devices (e.g., Van Dorn
sampler, Kemmerer bottle, plankton trap, pumps) is
then filtered through a net of appropriate size. In
weedy areas, a dip net or flexible collecting tube (Pennak, 1962) are very useful. Another simple method to
8. Phylum Rotifera
collect rotifers is to submerge a 3 – 4 L (1 gallon) glass
jar in a weedy region and to arrange loosely a few
aquatic plants in it. If using this technique, fill the jar to
about 2 – 5 cm from the top and place it near a subdued light source, such as a north window. Rotifers
which swim to the surface on the lighted side may be
removed using a transfer pipette.
Certain aquatic plants, such as Elodea, Myriophyllum, and filamentous algae are good substrata to examine for the presence of sessile rotifers, but Utricularia
usually provides the richest diversity (Wallace, 1980).
Plants with highly dissected leaves may be examined in
small dishes using a dissecting microscope. Broadleaved plants must be cut into strips and examined on
edge.
The upper few centimeters of moist sand taken just
above the water line along a lake or a marine shoreline
or the hyporheic interstitial zone of a stream bed usually provides several species of rotifers. Unfortunately,
very few studies have been conducted on rotifers from
this habitat termed “psammon,” in part because of the
difficulty in separating the organisms from the sand.
Recent studies have revealed a remarkable diversity
(35 – 85% of the fauna) and abundance (up to 105 individuals/L) of rotifers in this habitat. Under certain conditions rotifers may be found at depths of up to 60 cm
into the interstitial. For a review of these topics consult
Schmid-Araya (1995).
Sediments collected from the bottom using a
dredge, coring apparatus, or suction device usually
provide several bdelloid and monogonont species. The
upper few centimeters of sediments from a core collected in the winter or early spring normally contain
resting eggs, which can be induced to hatch within a
few days when several milliliters of sediment are incubated at ambient spring or summer temperatures (May,
1986, 1987).
Do not overlook laboratory aquaria or holding
tanks as potential sources of material. We have found
some unusual species in aquaria that had remained almost unattended for months. One might try adding a
small amount of sediments from several sites as a way of
adding variety to the rotifer community within the
aquarium. Sessile rotifers may be present if the aquarium
contains aquatic plants recently collected from the field.
However, if you attempt to keep sessile forms, be sure
first to remove all snails from the aquarium. Populations
of rotifers may be maintained or even increased by
adding a small amount of food once or twice a week.
B. Culture
King and Snell (1977), Ricci (1984), and Stemberger (1981) have summarized the general procedures
231
for culturing rotifers. Some species of rotifers may be
cultured in the laboratory quite easily, obtaining densities of 105 individuals/L in a few weeks. Others seem
impossible to keep even for short periods. Several
species are commonly raised under a variety of conditions (Table II). Most of these need regular care a few
times per week (changing the medium, feeding, etc.).
However, some species require little care. For example,
Habrotrocha rosa is easily cultured in dilute broths of
decaying powdered baby food and may be nearly ignored for weeks at a time without extinction of the culture (see Bateman, 1987). Techniques used to culture
protozoans (e.g., making extracts and infusions of various grains, manure, soil, etc.) have been adopted for
the culture of some rotifer species with great success.
Most rotifer cultures can be maintained xenically (i.e.,
containing several contaminating organisms) without
much problem. In fact, sometimes rotifers are found
contaminating cultures of protozoans, microcrustaceans, etc., that have been provided by commerical
biological supply companies. Rotifers have been maintained under axenic (Dougherty, 1963) or monoxenic
culture (Gilbert, 1970) conditions. Sophisticated culture techniques using single- and two-stage chemostats
have been described by several workers (Starkweather,
1987; Walz, 1993).
C. Preparation for Identification
Three preservatives are commonly used to preserve
rotifers: formalin and Lugol’s iodine at concentrations
of 5% or less, and ethanol at about 30 – 50%. Lugol’s
has two advantages over formalin. It is less toxic and it
stains the specimens slightly; the staining makes the animals more visible during sorting procedures. Unfortunately, Lugol’s iodine makes the lorica skrink and deforms the specimens. Other stains such as rose bengal
are commonly used to stain specimens preserved in either formalin or ethanol.
Unless special precautions are taken before fixation, illoricate rotifers (especially bdelloids) will contract into a completely unidentifiable lump, making
identification difficult if not impossible. Such specimens
have lost all value for taxonomic purposes, although
the trophi still may be useful (see later discussion). Histological fixatives (e.g., Bouins) or sugar-formalin solutions (used to prevent osmotic shock in microcrustaceans) are not particularly helpful in preventing
contraction in rotifers. Whenever possible, live specimens should be identified first, then fixed to determine
the effect of a particular fixative on body shape.
Fortunately, several anesthetics have been found to
work well on rotifers (Edmondson, 1959; May, 1985;
Nogrady and Keshmirian, 1986a, b; Pennak, 1989 p.
232
Robert Lee Wallace and Terry W. Snell
TABLE II
Examples of Rotifers That Have Been Cultured
Species
Bdelloids
Habrotrocha rosa
Philodina acuticornis
several bdelloids
Monogononts
Asplanchna spp.
Brachionus angularis
B. calyciflorus
B. patulus
B. plicatilis
B. rubens
Cephalodella forficula
Encentrum linnhei
Euchlanis dilatata
Keratella cochlearis
Keratella testudo
Lecane inermis
Notommata copeus
Polyarthra major
Polyarthra vulgaris
Synchaeta cecelia
Culture conditions
Reference
Batch cultures of tap water and
powdered baby food, dog food, etc.
Monoxenic culture
Batch cultures
Bateman (1987), Schramm and Becker (1987),
Wallace personal observations
Meadow and Barrow (1971)
Ricci (1984), Pourriot (1958)
Batch cultures
Aloia and Moretti (1973), Gilbert (1967),
Stemberger (1981), Stemberger and Gilbert (1984a)
Walz (1983a)
Gilbert (1963a)
Gilbert (1970)
Boraas and Bennet (1988)
Rao and Sarma (1986)
Hirayama and Ogawa (1972)
Lubzens (1987)
Droop and Scott (1978)
Pilarska (1972)
Schlüter and Groeneweg (1981)
Rothhaupt (1985)
Dodson (1984)
Scott (1983), Scott (1988)
King (1967)
Walz (1983a,b)
Stemberger and Gilbert (1987b)
Dougherty (1963)
Pourriot (1977)
Stemberger (1981)
Buikema et al. (1977)
Egloff (1988)
Continuous culture
Batch culture
Monoxenic cultures
Chemostat culture
Batch culture
Batch culture
Mass culture
Chemostat culture
Batch culture
Mass culture
Chemostat culture
Batch culture
Monoxenic and Chemostat cultures
Batch culture
Continuous and batch cultures
Batch culture
Batch culture
Batch culture
Batch culture
Batch culture
Batch culture
(For additional references see Hydrobiologia volumes 255/256 and 313/314; Pourriot (1965: 122) provides a list of species that have been
cultured prior to about 1959.)
195), but no single anesthetic has been shown to be
universally effective. Care should be taken when working with anesthetic agents; some are controlled substances and/or are toxic. An alternative approach is to
use the simple hot-water fixiation technique described
by Edmondson (1959, p. 433) which generally gives
excellent results with a number of species. It is generally a good idea to place live samples in jars over ice
for the return trip to the laboratory, although we have
found that a few species suffer when cooled (e.g.,
Sinantherina socialis).
Techniques for preservation and mounting of rotifers for examination using light microscopy (Edmondson, 1959, p. 433 Pennak, 1989, p. 196; De Smet, 1998)
and transmission and scanning electron microscopy
(Amsellem and Clément, 1980; Melone, 1998; Kleinow,
1998) have been described. However, the striking beauty
of the living rotifer is utterly lost during almost any fixation and mounting procedure.
Observations of live rotifers are not always easily
accomplished. The goal is to retard their movements
without crushing or distorting the specimen with the
cover glass. Both objectives may be accomplished with
a compression microscope slide (Taylor, 1993). If a
compressor is unavailable, then tiny pieces of broken
cover glass (not recommended) or clay supports (recommended) work well as corner supports. If the clay supports are too high, a slight pressure from a pencil on
each corner will reduce the height of the cover glass to
the desired level. With some practice, one can trap a
planktonic rotifer sufficiently to prevent swimming
without undue constriction. Sessile rotifers are handled
more easily. Plant material with attached rotifers can be
trimmed with iridectomy scissors to a size suitable for
placement on a microscope slide. The animal will remain in place without the need of compression as long
as the plant material is large enough to act as an anchor.
Methylcellulose or other viscous agents and fibrous
material, such as glass wool or shredded filter paper,
also may be used to impede swimming species. Unfortunately, methylcellulose interferes with ciliary function,
and fibers reduce observation to a game of hide-and-goseek. Any lighting conditions may be used, as long as
you are careful not to overheat the specimens. Strobe
lighting provides a marvelous view of ciliary movements, and darkfield illumination is often spectacular!
8. Phylum Rotifera
The final identification of many rotifer species requires examination of the trophi which, in certain
species, may be done by compressing an intact animal.
However, Myers (1937) described a method to extract
trophi from surrounding soft tissues using a small volume of bleach (sodium hypochlorite) in a depression
slide (see also Edmondson, 1959, pp. 432 – 433, Pennak,
1989, pp. 195 – 196; Stemberger, 1979). A regular microscope slide may be used, if the cover glass is
supported so the arrangements of the pieces of the
trophi are not disturbed by compression. Because the
trophi are liberated rather quickly when the bleach
comes into contact with the rotifer, it is necessary to find
the animal rapidly; otherwise it becomes necessary to
scan the entire slide for the small trophi. Be aware that
bubbles may form and obscure your view when bleach
comes into contact with some biological materials.
Methods for preparing trophi for scanning electron
microscope (SEM) work have been described by several
workers including a recent methodology by Kleinow
(1998) that gives stereoscopic pictures. In all work with
trophi, it is important to remember that these structures
are very small (50 m long) and that they are three
dimensional objects with a particular spatial arrangement among all seven pieces comprising the trophi.
V. CLASSIFICATION AND SYSTEMATICS
A. Classification
Classification schemes differ slightly in the ways
that they treat the three basic groups of rotifers. In this
key, three classes (Seisonidea, Bdelloidea, Monogononta) are recognized. The first two are comprised
of rotifers with two gonads and are sometimes considered orders within the class Digononta, leaving
class Monogononta separate (e.g., Pennak, 1989
p. 196). Another method of classification emphasizes
the unique anatomy and obligatory sexual reproduction of the Seisonidea, separating it from Bdelloidea
and Monogononta, which are then placed in the class
Eurotatoria (Koste, 1978, p. 48; Melone et al., 1998b).
European workers tend to rank rotifers as a class
within the phylum Aschelminthes, while North American authors treat rotifers as a phylum (in which case
the group known as aschelminths does not receive
taxon status).
B. Systematics
1. Class Seisonidea
This monogeneric taxon (Seison) comprises only
two recognized species of large (2 – 3 mm) dioecious,
233
marine rotifers that are epizoic on the gills of a leptostracan crustacean, Nebalia. The corona is very reduced and is not used in food gathering or locomotion.
Both species have paired gonads and a functional gut
in both sexes. The trophi are fulcrate. Females have
ovaries without vitellaria. Sexes are of similar size and
morphology. These unusual marine rotifers are not included in the following key. For a detailed review of
this class consult the work of Ricci et al., (1993).
2. Class Bdelloidea
Comprising 18 genera and just over 360 species
(Ricci 1987), bdelloids possess a rather uniform morphology and are exclusively females, reproducing by
parthenogenesis. Bdelloids are characterized by having
paired ovaries with vitellaria, more than two pedal
glands, and ramate trophi. Most bdelloids are microphagous with a corona of either two trochal disks
or a modified ciliated field. Bdelloids often have a vermiform body with a pseudosegmentation consisting of
annuli, which permit shortening and lengthening of the
body by telescoping. One order (Bdelloida) comprising
four families (Adinetidae, Habrotrochidae, Philodinavidae, Philodinidae) is recognized. Bdelloids generally are
not caught in plankton tows, although they may be
found in waters with dense vegetation. Bdelloids often
occur in sediments or among plant debris or crawling
on the surfaces of aquatic plants. Some forms inhabit
the capillary water films formed in soils (Pourriot
1979) or covering mosses. Many species are capable of
becoming desiccated and then rehydrated (see Section
II.C.4). Unfortunately, there has been no systematic review of North American bdelloids. The works of Bartos (1951), Burger (1948), Donner (1965), Koste and
Shiel (1986), and Pourriot (1979) should be consulted
for more detail on bdelloids.
3. Class Monogononta
Class Monogononta comprises the largest group of
rotifers with more than 1600 species in about 95 genera of benthic, free swimming, and sessile forms. All
are assumed to be dioecious with one gonad. In many
species, males have never been observed. Females possess one ovary with a vitellarium. Males usually are
structurally reduced with a vestigal gut that functions
only in energy storage. Males generally are ephemeral,
usually being present in the plankton for only a
few days or weeks each year. Monogononts are microphagous or raptorial, but a few are parasitic. The
corona is modified as broad-to-narrow disks or earlike
lobes or is vase-shaped with reduced ciliation and long
setae used in prey capture. In this key, three orders are
recognized (Collothecacea, Flosculariacea, Ploimida),
collectively comprising 24 families. Other workers
234
Robert Lee Wallace and Terry W. Snell
TABLE III List of the Rotifer Families Recognized in the Key
Presented Here
Order Bdelloidea
Adinetidae (4’)
Habrotrochidae (2)
Philodinavidae (4)
Philodinidae (3)
Order Collothecacea
Collothecidae (5)
Order Flosculariacea
Conochilidae (8)
Filiniidae (11)
Flosculariidae (9)
Hexarthridae (7)
Testudinellidae (11’)
Trochosphaeridae (10)
Order Ploimida
Asplanchnidae (15)
Birgeidae (14’)
Brachionidae (23’)
Colurellidae (19)
Dicranophoridae (12) [De Smet, 1997]
Epiphanidae (22)
Euchlanidae (21)
Gastropidae (27’)
Lecanidae (21’) [Segers, 1995a]
Lindiidae (14)
Microcodonidae (26)
Mytilinidae (20)
Notommatidae (26’) [Nogrady and Pourriot, 1995]
includes Ituridae [Pourriot, 1997] and Scaridiidae [Segers,
1995b]
Proalidae (22’) [De Smet, 1996]
Synchaetidae (27)
Trichocercidae (24)
Trichotriidae (23)
References in square brackets indicate recent publications done as a
part of an ongoing series of guides that are updating rotifer taxonomy.
include the first two orders as suborders within order
Gnesiotrocha (de Beauchamp, 1965; Koste, 1978;
Koste and Shiel, 1987). Table III lists the families recognized in the subsequent key and their relevant couplet number.
C. Taxonomic Keys
There are a variety of keys to rotifers including
those by Edmondson (1959) and Pennak (1989), covering the North American fauna, that may be followed to
the level of genus. However, the specialized keys of
Koste and Shiel (1987) for Australian waters, Pontin
(1978) for the British Isles, Stemberger (1979) for the
Great lakes, Ruttner-Kolisko (1974) for planktonic rotifers in general, and Koste’s (1978) revision of Voigt
for the rotifers of central Europe are also important.
This second group of works deals nearly exclusively
with monogonont rotifers. While these keys may not be
comprehensive enough to cover all of the variations in
size and morphology that are sometimes found within
a species, they will probably prove to be more than adequate for most work where detailed descriptions are
needed. A comprehensive work of providing a worldwide guide to the rotifers is currently underway under
the auspices of H.J.F. Dumont as coordinating editor.
Volume 1 in the series (Nogrady et al., 1993) provides
a general key to the level of family. To date four subsequent volumes have been published by SPB Academic
Publishers. These volumes collectively cover the following taxa: Volume 2, Lecanidae; Volume 3, Notommatidae and Scaridiidae; Volume 4, Proalidae; and Volume
5. Dicranophoridae. A very simple key is available
from Ward’s Natural Science.
One of the fundamental differences among the
higher taxonomic levels in phylum Rotifera is the structure of the trophi, with characteristic types of jaws being recognized in each family (see Section II.B.2). [Transitional forms are also known (Koste, 1978; Koste and
Shiel, 1987).] Although the following key is designed
with the nonspecialist in mind, commensurate with the
central importance of the trophi, we have used both the
structure of the trophi and other obvious characters of
anatomy and morphology as priniciple points of separation. In keeping with the nature of this text, the key is
taken only to the level of family following, for the
monogononts, the recent taxonomy of Koste (1978),
Koste and Shiel (1987), and Nogrady et al. (1993).
D. Taxonomic Key to Families of Freshwater Rotifera
1a.
Rotifers with paired ovaries (Digononta) and ramate trophi (Fig. 9) .......................................................................order Bdelloidea 2
1b.
Rotifers with a single ovary and trophi other than ramate ..................................................................................................................
...........................................................................................................................................................................class Monogononta 5
[Although this step obviously is very important, special care is generally not necessary to resolve this couplet. The ramate trophi of
bdelloids are usually identifable in whole animals without resorting to their isolation (see Melone et al., 1998b and Section IV.C). If
the type of trophi and number of ovaries cannot be determined, there are other clues that may be useful for live organisms. Upon
contacting a substrate, many bdelloids will crawl on the surface in a manner reminiscent of a leech, hence the etymon of the name
(Greek, bdella, leech). Further, the corona of some bdelloids has the appearance of two separate wheels, while only a few sessile
monogononts of the family Flosculariidae give this impression.]
8. Phylum Rotifera
FIGURE 37 Representatives of Family Habrotrochidae (Habrotrocha). (From Koste, 1976, with permission.)
235
FIGURE 38
Representatives of the Family Philodinidae. (From
Koste, 1976, with permission.)
2a(1a)
Stomach without lumen, as a syncitial mass of food vacuoles that gives the gut a frothy appearance ...................................................
.........................................................................................................................................................................family Habrotrochidae
[Three genera (e.g., Habrotrocha) reported in North America; mainly on moss or benthic, includes about 120 species (e.g.,
Fig. 37.]
2b
Stomach with lumen...........................................................................................................................................................................3
3a2b.
Corona with the appearance of two separate wheels (trochus) extended on pedicels..............................................family Philodinidae
[About 10 genera and 150 species, of which Philodina and Rotaria are common; also Dissotrocha and Macrotrachella
(Fig. 38).]
3b
Corona not as above ..........................................................................................................................................................................4
4a(3b).
Corona ciliated lobes or regions near mouth; rostrum ciliated............................................................................family Philodinavidae
[One rare species reported from North America (Abrochta, Fig. 39).]
4b.
Corona as flat ciliated fields on ventral side (no cingulum) .......................................................................................family Adinetidae
[Two genera (Adineta, Bradyscela) of about 15 species (Fig. 39).]
5a(1b).
Trophi uncinate (Fig. 8H); corona lacking typical ciliated regions, but is open as an infundibulum with or without elongate setae
around the margin ..............................................................order Collothecacea comprising 2 families: Atrochidae and Collothecidae
[Collothecidae (Collotheca and Stephanocerous) possess elongate setae (Figs. 6 and 40); fewer than 50 species; mostly sessile, ca.
five species are free swimming, many produce clear gelatinous tubes; none are colonial, but may colonize substrata forming dense
groupings (cf. Flosculariidae). Atrochidae (Acyclus, Atrochus, Cupelopagis) lack setae; each genus monospecific (Fig. 41).]
5b.
Trophi other than uncinate; corona with typical ciliated lobes or fields, or ciliated bands ..................................................................6
6a(5b).
Rotifers possessing malleoramate trophi (Fig.8I), order Flosculariacea...............................................................................................7
[Malleoramate trophi possess unci with numerous fine teeth nearly completely overlying the rami (unlike the malleate type; see the
description in couplet 16); teeth close to the fulcrum are usually larger than those more distant. In live animals, the nearly constant
movement of the trophi in a grinding or poundinglike action is characteristic.]
236
Robert Lee Wallace and Terry W. Snell
FIGURE 39 Representatives of two bdelloid families. (A) Family
Philodinavidae, Abrochta, ventral view of corona (adapted from
several sources); and (B) family Adinetidae, Adineta, dorsal view of
adult (from Koste, 1976, with permission).
FIGURE 40 Representative of Family Collothecidae (Collotheca
sp. within its gelatinous tube which is embedded with detritus and
diatoms).
6b
Rotifers with trophi other than malleoramate .........................................................................................................order Ploimida 12
7a(6a).
Conical body with six hollow, armlike, setose appendages that are outgrowths of the integument; appendages inserted with powerful muscles ............................................................................................................................................................family Hexarthridae
[Monogeneric family, genus Hexarthra (Pedalia) with about eight species (Fig. 13), some of which inhabit salt or brackish waters.
Although illoricate, Hexarthra spp. preserve well in formalin without serious contraction.]
7b.
Body lacking hollow, armlike, setose appendages ...............................................................................................................................8
8a(7b).
Rotifers with a corona of horseshoe or U-shaped ciliated bands; possessing a ventral gap in the coronal ciliation; free-swimming;
solitary or small to very large colonies (150 individuals) within a gelatinous matrix..........................................family Conochilidae
FIGURE 41
Representatives of family Atrochidae (Cupelopagis vorax). (Adapted from several sources.)
8. Phylum Rotifera
237
FIGURE 42 Coronae of the family Flosculariidae. (Modified from
several sources.)
[Two genera recognized, based on the position of antennae (Conochilus and Conochiloides), but Koste (1978) and Ruttner-Kolisko
(1974) do not consider antennal position to be a significant feature and subsume Conochiloides within Conochilus (Fig. 17).]
8a.
Rotifers with corona other than a horseshoe or U-shaped ciliated band; solitary or colonial; with or without a gelatinous
matrix ................................................................................................................................................................................................9
9a(8b).
Rotifers typically with elongate bodies and large, circular to lobate, earlike corona (Fig. 42); solitary or colonial; with or without a
tube or gelatinous matrix; mostly sessile, but some free swimming .....................................................................family Flosculariidae
[Seven genera of mainly sessile species (Figs. 5, 17C, and 43), a few are free-swimming; includes the genera Floscularia, Lacinularia,
Ptygura, and Sinantherina.]
9b.
Rotifers lacking a large, circular to lobate, earlike corona ...............................................................................................................10
10a(9b).
Spherical rotifers with a corona as a circular band around the equator or toward one end............................family Trochosphaeridae
[Genus Trochosphaera (Fig. 44) with two species in eutrophic waters, rare, but when present may be very abundant. A second rare
genus (Horaëlla) apparently not reported in the United States.]
10b.
Rotifers not spherical in shape .........................................................................................................................................................11
11a(10b).
Rotifers with two movable anterior spines (bristles) below the corona, of varying lengths often much longer than the body and one
(rarely two) rigid caudal spines; foot absent ..............................................................................................................family Filiniidae
[Genus Filinia (Fig. 32) with two North American species and apparently many hybrids. The long spines point posteriorly when the
animal is swimming, but the anterior ones point forward when the corona is withdrawn.]
11b.
Rotifers without long spines as described above ...............................................................................................family Testudinellidae
[Two dissimilar genera. (1) Genus Testudinella, lorica greatly flattened dorsoventrally, with dorsal and ventral plates fused along the
lateral margin; foot annulated and retractile ending in a ciliated cup, which is difficult to see; approximately 12 littoral species up to
250 m long (Fig. 45). (2) Genus Pompholyx, body having 4 nearly equal lobes in cross section; with a pair of frontal eyespots,
(Fig. 45) common in lakes and ponds.]
12a(6b).
Rotifers with forcipate trophi (Fig. 8J) ...........................................................................................................family Dicranophoridae
[About 12 genera of creeping, littoral rotifers (Fig. 46) with symmetrical or asymmetrical forcipate trophi, a feature easily determined by gentle compression of the specimen; no planktonic or semiplanktonic species are present in this family.]
12b.
Rotifers with trophi other than forcipate .........................................................................................................................................13
13a(12b).
Mostly littoral rotifers possessing cardate trophi (Fig. 10) having a sucking action or trophi highly modified and stomach with
zoochlorellae ...................................................................................................................................................................................14
13b.
Rotifers with trophi other than cardate ...........................................................................................................................................15
14a(13a).
Manubria of the trophi with hooks that may be determined in lateral view of the animal (Fig. 10); body spindle-shaped....................
family Lindiidae
[Monogenetic family, Genus Lindia (Fig. 47); mostly littoral.]
14b.
Highly modified trophi, possessing pseudunci; stomach with zoochlorellae, gastric glands absent (Fig. 48) ................family Birgeidae
[Monospecific family (Birgea enantia, a rare littoral species).]
FIGURE 43 Representatives of family Flosculariidae. (A) Floscularia conifera, portion of a sessile colony, with two
adults inside a tube constructed of pellets made of detritus (tube length may reach 2000 m); (B) Ptygura barbata attached
to a filamentous alga (total animal length ⬇ 300 m); (C) Limnias melicerta, sessile in its clear cement tube (individuals ⬇
900 m); and (D) Ptygura mucicola, sessile within the colonial cyanobacterium, Gloeotrichia (body length ⬇ 100 m).
238
8. Phylum Rotifera
239
FIGURE 44
Representative of family Trochosphaeridae. (A) Photomicrograph of Trochosphaera (specimen slightly distorted); and (B)
line drawing interpretation. (Diameter of the individuals ⬇ 800 m; original photomicrograph by Tom Nogrady.)
15a(13b).
Saccate rotifers, with incudate or modified incudate trophi, (Fig. 8G,R) some lacking intestine and anus...........family Asplanchnidae
[Three genera. Two lacking an intestine: Asplanchna, foot absent, with six species (Figs. 11 and 49) and Asplanchnopus, foot present, with three species. One rare, benthic genus with intestine and foot, Harringia, (Fig. 50) with two species.]
15b.
Rotifers not possessing incudate trophi ............................................................................................................................................16
16a(15b).
Rotifers possessing malleate trophi...................................................................................................................................................17
[In malleate trophi (Fig. 8A,B,C,E,F,P), each uncus has only 4 – 7 teeth (unlike the condition found in malleoramate trophi; see description couplet 6). The fulcrum may be short (malleate) or long (submalleate).]
16b.
Rotifers possessing virgate trophi ....................................................................................................................................................24
[Virgate trophi are often asymmetrical (Fig. 8D,K,L,M,N,O,Q,S), with a long fulcrum and manubria and generally small rami.]
FIGURE 45 Representatives of family Testudinellidae. (Ventral
views of Testudinella and Pompholyx.) (Modified from several
sources.)
FIGURE 46 Representative of family Dicranophoridae (Dicranophorus). (From Koste, 1976, with permission.)
240
Robert Lee Wallace and Terry W. Snell
FIGURE 47
Representative of family Lindiidae (Lindia). (From
Harring and Myers, 1922, with permission.)
FIGURE 48 Representative of family Birgeidae (Birgea). (From
Harring and Myers, 1922, with permission.)
17a(16a).
Loricate rotifers: body wall thickened and firm ................................................................................................................................18
17b.
Illoricate rotifers: body wall not thickened or firm ...........................................................................................................................22
[Interpreting whether the body wall is thickened (loricate) or not (illoricate) can be difficult and it does take some experience to
judge this characteristic. To get an indication as to how firm the lorica is, follow this procedure. When working with fresh material gently, apply pressure from the point of a pencil or probe onto the cover glass while observing the specimen. If the body
puffs out under pressure and returns to its original shape when the pressure is released, the specimen is illoricate. Loricate forms
will exhibit much less flexibility of the body wall. In preserved materials, illoricate forms tend to shrivel up, while the body
wall of loricate rotifers will retain its shape even if the inner organs separate from the body wall, collapsing into a central mass
of tissue.]
18a(17a).
Lorica possessing furrows, grooves, or sulci; or with a dorsal, semicircular head shield covering the corona; or with a very strongly
developed lorica and a dorsal transverse ridge..................................................................................................................................19
18b.
Lorica lacking furrows, grooves, sulci or dorsal head shield; without a strongly developed lorica and dorsal transverse ridge ........23
FIGURE 49 Representative of family Asplanchnidae (Asplanchna).
[From Koste, 1976, with permission.]
FIGURE 50
Representative of family Asplanchnidae (Harringia).
[From Koste, 1976, with permission.]
8. Phylum Rotifera
FIGURE 51 Representative of family Colurellidae. [Modified from
several sources.]
241
FIGURE 52
Representative of family Mytilinidae (Mytilina). [Modified from several sources.]
19a(18a).
Lorica with medial, ventral furrow (but no lateral furrow or grooves) extending the full length of the animal or with a ventral notch
in which the foot lies, or with a dorsal, semicircular head shield covering the corona ............................................family Colurellidae
[Four genera found in the littoral zone including Colurella (possessing a medial ventral furrow, Fig. 51A), a very common genus of
some 15 species, which browse on epiphytic microorganisms; Lepadella (possessing a ventral notch in which the foot lies, Fig. 51B),
very common with many species; Squatinella (possessing a dorsal semicircular head shield covering the corona, Fig. 51C). [N.B.:
Diplois daviesiae (family Euchlanidae), a rare monospecific genus present in Sphagnum bogs will key to family Colurellidae, if the
lateral furrows of this species are missed.]
19b.
Lorica lacking medial, ventral furrow or notch, and lacking a head shield .......................................................................................20
20a(19b).
Lorica with dorsal, medial sulcus (double keel) or with a strongly developed lorica usually possessing a dorsal transverse ridge ........
family Mytilinidae
[Two littoral genera: Mytilina (Fig. 52) possessing a laterally flattened lorica with a dorsal longitidutinal sulcus, spines on all four
anterior corners, about 10 species common, and Lophocharis with a strongly developed lorica, usually having a dorsal transverse
ridge.]
20b.
Lorica lacking a dorsal, medial sulcus and lacking a dorsal transverse ridge ....................................................................................21
▼
FIGURE 53 Representative of family Euchlanidae (Euchlanis).
[Modified from several sources.]
242
Robert Lee Wallace and Terry W. Snell
▼
FIGURE 54
Representatives of the family Lecanidae. (A & B)
Lecane spp.; (C) Lecane lunaris (dorsal); and (D) lateral view. (A and
B from several sources; C and D from Dartnall and Hollowday, 1985,
with permission.)
21a(20b)
Foot projecting from between dorsal and ventral plates at the posterior end of the lorica; dorsal and ventral plates separated by a
deep furrow or groove ...........................................................................................................................................family Euchlanidae
[Four genera common in the littoral rotifers, including Euchlanis (Fig. 53).]
21b.
Foot projecting through a hole in the ventral plate at the posterior end of the lorica; dorsal and ventral plates connected by a weak
furrow or groove ......................................................................................................................................................family Lecanidae
▼
FIGURE 55 Representative of family Epiphanidae (Epiphanes).
(From Dartnall and Hollowday, 1985, with permission.)
8. Phylum Rotifera
243
FIGURE 56 Representatives of family Proalidae. (A) Proales (from Chengalath, 1985, with permission); and
(B) photomicrograph of adult of Proales werneckii (dark body) with numerous eggs within a gall of Vaucheria
sp. [Gall ⬇ 600 m long.]
[A large number of littoral species in two genera (Fig. 54): Lecane (with two toes which may be partially fused at the base) and
Monostyla (one toe or toe divided distally); Koste (1978) has subsummed Monostyla within Lecane.]
22a(17b).
Mouth set in a funnel-shaped buccal field ..............................................................................................................family Epiphanidae
[Six genera including Epiphanes (Fig. 55), Cyrtonia, Mikrocodides, and Rhinoglena all littoral species or in small, shallow lakes.
Epiphanes, sometimes incorrectly described as a ‘typical rotifer’ in textbooks, is not common, but may be found in ponds receiving
animal wastes.]
22b.
Mouth set in an oblique, ciliated field on ventral side; body wormlike to fusiform ....................................................family Proalidae
[Four genera, including the large genus Proales; generally found in littoral zones and sandy beaches (Fig. 56A). Proales daphnicola
is planktonic and may be attached to cladocerans; Proales werneckii is parasitic in galls that it induces in filaments of aquatic
Vaucheria spp (Fig. 56B).]
23a(18b).
Lorica extending beyond body to head, foot, and toes ..........................................................................................family Trichotriidae
[Three genera, including Macrochaetus (Fig. 57A), with numerous bilaterally placed spines (about seven littoral species) and Trichotria (Fig. 57B), with a pair of heavy spines on the dorsal side of the foot (about 10 littoral species).]
23b.
Lorica not extending beyond body .......................................................................................................................Family Brachionidae
[A large family comprising 6 genera of common rotifers. Brachionus, a very common genus with about 25 species of littoral and
planktonic rotifers (Fig. 58). B. plicatilis is frequently found in salt and brackish waters. Kellicottia, with two common planktonic
species, possessing long anterior spines of unequal length (Fig. 59). Keratella, with more than 15 species, having the dorsal
244
Robert Lee Wallace and Terry W. Snell
▼
FIGURE 57
Representatives of family Trichotriidae (Macrochaetus
and Trichotria). [Modified from several sources.]
plate decorated with a characteristic facet pattern (Fig. 60). K. cochlearis is very common. Notholca a genus common in cool
waters (Fig. 61).]
24a(16b).
Body twisted as a partial helix (asymmetrical) and/or trophi asymmetrical; or small, saccate animals, predatory within the colonial
alga, Volvox .......................................................................................................................................................family Trichocercidae
[Three genera: (1) genus Trichocera (body twisted as a partical helix; unequal toes; asymmetrical trophi) with some 90 species,
may be important in the plankton (Fig. 62); (2) genus Elosa (body showing in cross section three lobes, two lateral and one dorsal,
plus ventrally an inconspicous lobe; asymmetrical trophi) with two species common in the psammon and with Sphagnum moss;
(3) Ascomorphella volvocicola, predatory within Volvox (Fig. 63).]
24b.
Body and trophi not as described above ...........................................................................................................................................25
25a(24b).
Free-swimming rotifers.....................................................................................................................................................................27
25b.
Crawling or creeping rotifers in the littoral (ocassional species in the plankton) ..............................................................................26
FIGURE 58
Representatives of brachionid rotifers (family Brachionidae; genus Brachionus. [From Koste, 1976, with permission.]
FIGURE 59 Kellicottia (dorsal view) a common genus of the family
Brachionidae. (From several sources.)
8. Phylum Rotifera
245
Variation in the genus Keratella, a common genus of the family Brachionidae. (Individuals ⬇
100 – 200 m in length.) (A) K. cochlearis (from Koste, 1976, with permission); (B) K. quadrata; and (C) K.
testudo (from Stemberger, 1988, with permission).
FIGURE 60
26a(25b).
Purple plates positioned anterior to the mastax; corona wide, flat, and somewhat circular; foot jointed and long, about 50% of the
total length of the animal, with a single toe ....................................................................................................family Microcodonidae
[Monospecific family of one uncommon species, Microcodon clavus, mostly littoral, but may be found in the plankton.]
26b.
Lacking purple plates as described earlier, corona ventral and not circular ........................................................family Notommatidae
[A varied family with many species in 13 genera including Cephalodella (Fig. 64), Notommata (Fig. 65), and Pleurotrocha. Also included at this point in the key are the families Ituridae and Scaridiidae.]
27a(25a).
Corona with four prominant setae (sensory bristles) and auricles (earlike structures); or possessing 12 movable, flattened, sword-topaddle- or feather-shaped appendages (paddles); or with a sculptured lorica having ridges, grooves, or areolations ............................
family Synchaetidae
[Four genera: with sensory bristles and auricles, Synchaeta (Fig. 66), comprising some 12 freshwater species plus others of marine
and brackish waters, may be important in the plankton; with paddles Polyarthra (Fig. 33), fewer than 10 species separated based
on paddle morphology, nuclei of vitellarium, and lateral antennae; Pseudoploesoma (Fig. 67A) and Ploesoma (Fig. 67B).]
FIGURE 61 Notholca, a common genus of the family Brachionidae, usually found in cool waters. (A) Variation within the genus (modified from several sources); and (B) N. squamula. (Modified from May, 1980, with
permission).
246
Robert Lee Wallace and Terry W. Snell
FIGURE 62 Trichocera (lateral view) an important genus of the
family Trichocercidae. Lateral view. [From Wallace et al., 1989, with
permission.]
FIGURE 63
Predatory activity of Ascomorphella volvocicola (family Trichocercidae) on its prey, the colonial
alga Volvox. Seen here are at least three rotifers feeding on the cells of a colony. Gaps in the cells of the alga
represent the damage done by the rotifer; an egg is seen just to the right of center. (Photomicrograph by courtesy
of Russ Shiel.)
8. Phylum Rotifera
FIGURE 64 Representatives of family Notommatidae (Cephalodella). (Cephalodella sp. from Dartnall and Hollowday, 1985;
others from Koste, 1976, with permission.)
27b.
247
FIGURE 65 Representative of family Notommatidae (Notommata,
dorsal view). (From Koste, 1976, with permission.)
Lacking prominant setae, auricles, and paddles, and lorica not sculptured .........................................................family Gastropodidae
[Two genera: Ascomorpha (Fig. 8.68A) (incorporating Chromogaster) comprising 6 species, possessing a fingerlike projection
(palp) from the corona; Gastropus (Fig. 8.68B) with three species having a laterally compressed body, may be important in the
plankton.]
FIGURE 66
Representatives of family Synchaetidae (Synchaeta).
[From Koste, 1976, with permission.]
FIGURE 67 Representatives of family Synchaetidae. (A) Pseudoploesoma (side view) and (B) Ploesoma (dorsal view). Bars ⬇ 75 m.
(Modified from several sources.)
248
Robert Lee Wallace and Terry W. Snell
Representatives of family Gastropodidae. (A) Ascomorpha sp.; (B) Gastropus, (bar 100 m).
(A from Koste, 1976, with permission; and B modified from several sources.)
FIGURE 68
ACKNOWLEDGMENTS
We wish to thank our mentors J. J. Gilbert and C.
E. King for all the help and encouragement they have
given us over the years. We also acknowledge Dr. W.
Koste, a good friend for advice and inspiration; a
worker who knows more about rotifers than we ever
thought possible. Drs. W. T. Edmondson, J. J. Gilbert,
T. Nogrady, P. L. Starkweather, R. S. Stemberger, L. J.
Walsh, and G. H. Wittler reviewed the manuscript for
this chapter, a task very much appreciated by the authors. However, we remain responsible for any errors
which may still be present. We thank H. L. “Chico”
Taylor for his insight in preserving and mounting rotifers for taxonomic work.
LITERATURE CITED
Allan, J. D. 1976. Life history patterns in zooplankton. American
Naturalist 110:165 – 180.
Aloia, R. C., Moretti, R. L. 1973. Sterile culture techniques for some
species of the rotifer Asplanchna. Transactions of the American
Microscopical Society 92:364 – 371.
Amsellem, J., Clément, P. 1980. A simplified method for the preparation of rotifers for transmission and scanning electron microscopy. Hydrobiology 73:119 – 122.
Amsellem, J., Ricci, C. 1982. Fine structure of the female genital apparatus of Philodina (Rotifera, Bdelloidea). Zoomorphology
100:89 – 105.
Amrén, H. 1964. Ecological and taxonomic studies on zooplankton
from Spitsbergen. Zool. Bidr. Uppsala 36:193 – 208.
Arndt, H. 1993. Rotifers as predators on components of the microbial web (bacteria, heterotrophic flagellates, ciliates) — a review.
Hydrobiologia 255/256:231 – 246.
ASTM (American Society of Testing and Materials). 1991. Standard
guide for acute toxicity test with the rotifer Brachionus. Vol.
11.05, E 1440, West Conshohocken, PA.
Barron, G. L. 1980a. Fungal parasites of rotifers: Harposporium.
Canadian Journal of Botany 58:443 – 446.
Barron, G. L. 1980b. A new Haptoglossa attacking rotifers by rapid
injection of an infective sporidium. Mycologia 72:1186 – 1194.
Barron, G. L., Tzean, S. S. 1981. A subcuticular endoparasite impaling bdelloid rotifers using three-pronged spores. Canadian Journal of Botany 59:1207 – 1212.
Bartos, E. 1951. The Czechoslovak Rotatoria of the order Bdelloidea.
Vestník Ceskoslovenské Zoologické Spolecnosti 15:241 – 500.
Bateman, L. 1987. A bdelloid rotifer living as an inquiline in leaves
of the pitcher plant, Sarracenia purpurea. Hydrobiologia 147:
129 – 133.
de Beauchamp, P. 1965. Classe des Rotifères, in: Grassé, P. P. Ed.,
Traité de Zoologie IV, 3, Masson, Paris, pp. 1225 – 1379.
Bender, K., Kleinow, W. 1988. Chemical properties of the lorica and
related parts from the integument of Brachionus plicatilis. Comparative Biochemistry and Physiology 89B:483 – 487.
Berzins, B., Pejler, B. 1987. Rotifer occurrence in relation to pH. Hydrobiologia 147:107 – 116.
Birky, Jr., C. W., 1967. Studies on the physiology and genetics of the
rotifer Asplanchna III. Results of outcrossing, selfing, and selection. Journal of Experimental Zoology 165:104 – 116.
Bogdan, K. G., Gilbert, J. J. 1982. Seasonal patterns of feeding by natural populations of Keratella, Polyarthra, and Bosmina: clearance
rates, selectivities, and contributions to community grazing. Limnology and Oceanography 27:918 – 934.
Bogdan, K. G., Gilbert, J. J. 1987. Quantitative comparison of food
niches in some freshwater zooplankton. Oecologia (Berlin) 72:
331 – 340.
Boraas, M. E., Bennett, W. N. 1988. Steady-state rotifer growth in a
two-stage, computer-controlled turbidostat. Journal of Plankton
Research 10:1023 – 1038.
Brett, M. T. 1989. Zooplankton communities and acidification
processes (a review). Water, Air and Soil Pollution 44:387 – 414.
Buikema, A. L. Jr., Cairns, Jr., J., Edmunds, P. C., Krakauer, T. H.
1977. Culturing and ecology studies of the rotifer, Polyarthra
8. Phylum Rotifera
vulgaris. EPA-600/3-77-051. Environmental Research Laboratory, Office of Research and Development, U.S. Environmental
Protection Agency, Duluth, Minnesota.
Burger, A. 1948. Studies on the moss dwelling bdelloids (Rotifera) of
eastern Massachusetts. Transactions of the American Microscopical Society 67:111 – 142.
Burns, C. W., Gilbert, J. J. 1986a. Effects of daphnid size and density
on interference between Daphnia and Keratella cochlearis. Limnology and Oceanography 31:848 – 858.
Burns, C. W., Gilbert, J. J. 1986b. Direct observations of the mechanisms of interference between Daphnia and Keratella cochlearis.
Limnology and Oceanography 31:859 – 866.
Capuzzo, J. 1979a. The effect of temperature on the toxicity of chlorinated cooling waters to marine animals — a preliminary review. Marine Pollution Bulletin 10:45 – 47.
Capuzzo, J. 1979b. The effect of halogen toxicants on survival, feeding and egg production of the rotifer Brachionus plicatilis. Estuarine and Coastal Marine Science 8:307 – 316.
Charoy, C. 1995. Modification of the swimming behaviour of Brachionus calyciflorus (Pallas) according to food environment and
individual nutritive state. Hydrobiologia 313/314: 197–204.
Chengalath, R., Koste, W. 1987. Rotifera from northwestern Canada.
Hydrobiologia 147:49 – 56.
Clément, P. 1977. Ultrastructural research on rotifers. Archiv für Hydrobiologie Beiheft 8:270 – 297.
Clément, P. 1980. Phylogenetic relationships of rotifers, as derived
from photoreceptor morphology and other ultrastructural analysis. Hydrobiologia 73:93 – 117.
Clément, P. 1985. The relationships of rotifers, in: Conway Morris,
S., George, J. D., Platt, H. M., Eds. The origins and relationships of lower invertebrates. Systematics Association, Vol. 28,
Clarendon Press, Oxford, pp. 224 – 247.
Clément, P. 1987. Movements in rotifers: correlations of ultrastructure and behavior. Hydrobiologia 147:339 – 359.
Clément, P., Wurdak, E., Amsellem, J. 1983. Behavior and ultrastructure of sensory organs in rotifers. Hydrobiologia 104:89 – 130.
Clément, P., Wurdak, E. 1991. Rotifera. in: Harrison, F. W., Ruppert,
E. E. Eds. Microscopic Anatomy of Invertebrates, Vol. 4: Aschelminthes, Wiley-Liss, New York, pp. 219 – 297.
Couillard, Y., Ross, P., Pinel-Alloul, B. 1989. Acute toxicity of six
metals to the rotifer Brachionus calyciflorus. Toxicity Assessment 4:451 – 462.
Coulon, P. Y., Charras, J. P., Chasse, J. L., Clément, P., Cornillac, A.,
Luciani, A., Wurdak, E. 1983. An experimental system for automatic tracking and analysis of rotifer swimming behavior. Hydrobiologia 104:197 – 202.
Dad, N. K., Kant Pandya, V. 1982. Acute toxicity of two insecticides
to rotifer Brachionus calyciflorus. International Journal of Environmental Studies 18:245 – 246.
Dartnall, H. J. G., Hollowday, E. D. 1985. Anarctic rotifers. British
Antarctic Survey Scientific Reports 100:1 – 46.
De Smet, W. 1996. in: Dumont, H. J. (Ed.) Rotifera, Vol. 4: The
Proalidae (Monogononta). Guide to the identification of the
microinvertebrates of the continental waters of the world. SPB
Academic Publishers, The Hague, The Netherlands. 102 pp.
De Smet, W. 1997. Rotifera in: Dumont, H. J., Vol. 5: The
Dicranophoridae (Monogononta). Guide to the identification
of the microinvertebrates of the continental waters of the
world. SPB Academic Publishers, The Hague, The Netherlands.
344 pp.
De Smet, W. 1998. Preparation of rotifer trophi for light microscopy.
Hydrobiologia 387/388:117 – 121.
Diéguez, M., Balseiro, E. 1998. Colony size in Conochilus hippocrepis: defense adaptation to predator stage sizes. Hydrobiologia
387/388:421 – 425.
249
Dodson, S. I. 1984. Ecology and behaviour of a free-swimming tubedwelling rotifer Cephalodella forficula. Freshwater Biology 14:
329 – 334.
Donner, J. 1965. Ordnung Bdelloidea. Bestimmungsbücher zur Bodenfauna Europas. Vol. 6. Akademie Verlag, Berlin. 267 pp.
Dougherty, E. C. 1963. Cultivation and nutrition of micrometazoa
III. The minute rotifer Lecane inermis. Journal of Experimental
Zoology 153:183 – 186.
Droop, M. R., Scott, J. M. 1978. Steady-state energetics of a planktonic herbivore. Journal of the Marine Biological Association of
the United Kingdom 58:749 – 772.
Dumont, H. 1983. Biogeography of rotifers. Hydrobiologia 104:
19 – 30.
Dumont, H., Segers, H. 1996. Estimating lacustrine zooplankton
species richness and complimentarity. Hydrobiologia 341:
125 – 132.
Edmondson, W. T. 1944. Ecological studies of sessile Rotatoria, Part I.
Factors affecting distribution. Ecological Monographs 14:32 – 66.
Edmondson, W. T. 1945. Ecological studies of sessile Rotatoria, Part
II. Dynamics of populations and social structure. Ecological
Monographs 15:141 – 172.
Edmondson, W. T. 1959. Rotifera, in: W. T. Edmondson, Ed. Freshwater Biology, 2nd Ed. Wiley, New York, pp. 420 – 494.
Edmondson, W. T. 1965. Reproductive rate of planktonic rotifers as
related to food and temperature in nature. Ecological Monographs 35:61 – 111.
Edmondson, W. T. 1993. Experiments and quasi-experiments in limnology. Bulletin of Marine Science 53:65 – 83.
Edmondson, W. T., Litt, A. H. 1982. Daphnia in Lake Washington.
Limnology and Oceanography 30:180 – 188.
Edmondson, W. T., Litt, A. H. 1987. Conochilus in Lake Washington. Hydrobiologia 147:157 – 162.
Egloff, D. A. 1988. Food and growth relations of the marine microzooplankter, Synchaeta cecelia (Rotifera). Hydrobiologia 157:
129 – 141.
Enesco, H. E. 1993. Rotifers in aging research: use of rotifers to test
various theories of aging. Hydrobiologia 255/256:59 – 70.
Enesco, H. E., Verdone-Smith, C. 1980. a-Tocopherol increases lifespan in the rotifer Philodina. Experimental Gerontology 15:
335 – 338.
Epp, R. W., Lewis, W. M. 1979. Sexual dimorphism in Brachionus
plicatilis (Rotifera): evolutionary and adaptive significance. Evolution 33:919 – 928.
Epp, R. W., Lewis, W. M. 1980. Metabolic uniformity over the environmental temperature range in Brachionus plicatilis (Rotifera).
Hydrobiologia 73:145 – 147.
Epp, R. W., Lewis, W. M. 1984. Cost and speed of locomotion for
rotifers. Oecologia 61:289 – 292.
Epp, R. W., Winston, P. W. 1977. Osmotic regulation in the brackishwater rotifer Brachionus plicatilis (Muller). Journal of Experimental Biology 68:151 – 156.
Epp, R. W., Winston, P. W. 1978. The effects of salinity and pH on the
activity and oxygen consumption of Brachionus plicatilis (Rotatoria). Comparative Biochemistry and Physiology 59A:9 – 12.
Felix, A., Stevens, M. E., Wallace, R. L. 1995. Unpalatability of a colonial rotifer, Sinantherina socialis, to small zooplanktivorous fishes.
Invertebr. Biol. 114:139 – 144.
Finlay, B. J., Curds, C. R., Bamforth, S. S., Bafort, J. M. 1987. Cilated
Protozoa and other microorganisms from two African soda
lakes (Lake Nakuru and Lake Simbi, Kenya). Arch für Protistenkunde 133:81 – 91.
Frost, T. M., Montz, P. K., Gonzalez, M. J., Sanderson, B., Arnott, S.
1998. Rotifer responses to increased acidity: long-term patterns
during the experimental and two manipulation of Little Rock
Lake. Hydrobiologia 387/388:141 – 152.
250
Robert Lee Wallace and Terry W. Snell
Garey, J. R., Near, T. J., Nonnemacher, M. R., Nadler, S. A. 1996.
Molecular evidence for Acanthocephala as a subtaxon of Rotifera. Journal of Molecular Evolution 43:287 – 292.
Garey, J. R., Schmidt-Rhaesa, A. Near, T. J. Nadler, S. A. 1998. The
evolutionary relationships of rotifers and acanthocephalans.
Hydrobiologia 387/388:83 – 91.
Gilbert, J. J. 1963a. Mictic female production in the rotifer Brachionus
calyciflorus. Journal of Experimental Zoology 153:113 – 123.
Gilbert, J. J. 1963b. Contact chemoreceptors, mating behavior and
reproductive isolation in the rotifer genus Brachionus. Journal
of Experimental Biology 40:625 – 641.
Gilbert, J. J. 1966. Rotifer ecology and embryological induction. Science 151:1234 – 1237.
Gilbert, J. J. 1967. Asplanchna and postero-lateral spine production
in Brachionus calyciflorus. Archiv für Hydrobiologie 64:1 – 62.
Gilbert, J. J. 1970. Monoxenic cultivation of the rotifer Brachionus
calyciflorus in a defined medium. Oecologia (Berlin) 4:89 – 101.
Gilbert, J. J. 1977. Effect of the non-tocopherol component of the diet
on polymorphism, sexuality and reproductive rate of the rotifer
Asplanchna sieboldi. Archiv für Hydrobiologie 80:375 – 397.
Gilbert, J. J. 1980a. Female polymorphisms and sexual reproduction in
the rotifer Asplanchna: evolution of their relationship and control
by dietary tocopherol. American Naturalist 116:409 – 431.
Gilbert, J. J. 1980b. Feeding in the rotifer Asplanchna: behavior, cannibalism, selectivity, prey defenses, and impact on rotifer communities, in: W. C. Kerfoot, Ed., Evolution and ecology of zooplankton communities. Univ. Press of New England, Hanover,
NH, pp. 158 – 172.
Gilbert, J. J. 1985a. Escape response of the rotifer Polyarthra: a highspeed cinematographic analysis. Oecologia (Berlin) 66:322 – 331.
Gilbert, J. J. 1985b. Competition between rotifers and Daphnia.
Ecology 66:1943 – 1950.
Gilbert, J. J. 1987. The Polyarthra escape response: defense against
interference from Daphnia. Hydrobiologia 147:235 – 238.
Gilbert, J. J. 1988. Suppression of rotifer populations by Daphnia: a
review of the evidence, the mechanisms, and the effects on zooplankton community structure. Limnology and Oceanography
33(6, part 1):1286 – 1303.
Gilbert, J. J. 1989a. The effect of Daphnia interference on a natural
rotifer and ciliate community: short – term bottle experiments.
Limnology and Oceanography 34(3):606 – 617.
Gilbert, J. J. 1989b. Competitive interactions between the rotifer
Synchaeta oblonga and the cladoceran Scapholeberis kingi Sars.
Hydrobiologia 186/187:75:80.
Gilbert, J. J. 1990. Differential effects of Anabena affinis on cladocerans and rotifers: mechanisms and implications. Ecology 71:
1727 – 1740.
Gilbert, J. J. 1995. Structure, development and induction of a new diapause stage in rotifers. Freshwater Biology 34:263 – 270.
Gilbert, J. J., Bogdan, K. G. 1984. Rotifer grazing: in situ studies on
selectivity and rates, in: Meyers, D. G., Strickler, J. R. Eds.,
Trophic Interactions within Aquatic Ecosystems. American Association for the Advancement of Science, Selected Symposium
85. Westview, Boulder, CO, USA pp. 97 – 133.
Gilbert, J. J., Jack, J. D. 1993. Rotifers as predators on ciliates. Hydrobiologia 255/256:247 – 253.
Gilbert, J. J., Kirk, K. L. 1988. Escape response of the rotifer Keratella:
description, stimulation, fluid dynamics, and ecological significance. Limnology and Oceanography 33(6, Part 2):1440 – 1450.
Gilbert, J. J., Starkweather, P. L. 1977. Feeding in the rotifer
Brachionus calyciflorus I. Regulatory mechanisms. Oecologia
(Berlin) 28:125 – 131.
Gilbert, J. J., Starkweather, P. L. 1978. Feeding in the rotifer Brachionus calyciflorus III. Direct observations on the effects of
food type, food density, change in food type, and starvation on
the incidence of pseudotrochal screening. Internationale Vereinigung für theoretische und angewandte Limnologie, Verhandlungen 20:2382 – 2388.
Gilbert, J. J., Stemberger, R. S. 1985a. Control of Keratella populations by interference competition from Daphnia. Limnology and
Oceanography 30:180 – 188.
Gilbert, J. J., Stemberger, R. S. 1985b. Prey capture in the rotifer
Asplanchna girodi. Internationale Vereinigung für theoretische
und angewandte Limnologie, Verhandlungen 22:2997 – 3000.
Gilbert, J. J., Stemberger, R. S. 1985c. The costs and benefits of gigantism in polymorphic species of the rotifer Asplanchna.
Archiv Für Hydrobiologie, Beiheft 21:185 – 192.
Gilbert, J. J., Williamson, C. E. 1978. Predator – prey behavior and
its effect on rotifer survival in associations of Mesocyclops
edax, Asplanchna girodi, Polyarthra vulgaris, and Keratella
cochlearis. Oecologia (Berlin) 37:13 – 22.
Gonzalez, M. J., Frost. T. M. 1994. Comparisons of laboratory
bioassays and a whole lake experiment: Rotifer responses to experimental acidification. Ecological Applications 4:69 – 80.
Green, J. 1972. Latitudinal variations in associations of planktonic
rotifers. Journal of Zoology, London 167:31 – 39.
Green, J. 1977. Dwarfing of rotifers in tropical crater lakes. Archiv
für Hydrobiologie Beiheft 8:232 – 236.
Green, J. 1985. Horizontal variations in associations of zooplankton
in Lake Kariba. Journal of Zoology, London 206:225 – 239.
Gvozdov, A. O. 1986. Phototaxis as a test function in bioassay.
Gidrobiologichesky Zhurnal 22:65 – 68.
Hagiwara, A., Snell, T. W., Lubzens, E., Tamaru, C. (Eds.). 1997.
Live food in Aquaculture., Vol. 358, Kluwer Academic Publishers, Belgium, Hydrobiologia.
Halbach, U. 1970. Die Ursachen der Temporal Variation von Brachionus calyciflorus Pallas (Rotatoria). Oecologia 4:262 – 318.
Halbach, U. 1984. Population dynamics of rotifers and its consequences for ecotoxicology. Hydrobiologia 109:79 – 96.
Halbach, U., Halbach – Keup, G. 1974. Quantitative Beiehungen
zwischen Phytoplankton und der Populationdynamik des Rotators Brachionus calyciflorus Pallas. Befunde aus Laboratoriums – experimenten und Freilanduntersuchungen. Archiv für
Hydrobiologie 73:273 – 309.
Halbach, U., Wiebert, M., Westermayer, M., Wissel, C. 1983. Population ecology of rotifers as a bioassay tool for ecotoxicological
tests in aquatic environments. Ecotoxicology and Environmental Safety 7:484 – 513.
Halbach, U., Wiebert, M., Wissel, C., Kalus, J., Beuter, K., Delion,
M. 1981. Population dynamics of rotifers as bioassay tools for
toxic effects of organic pollutants. Internationale Vereinigung
für theoretische und angewandte Limnologie, Verhandlungen
21:1141 – 1146.
Hall, D. J., Threlkeld, S. T., Burns, C. W., Crowley, P. H. 1976. The
size – efficiency hypothesis and the size structure of zooplankton
communities. Annual Review of Ecology and Systematics 7:
177 – 208.
Harring, H. K., Myers, F. J. 1922. The rotifers of Wisconsin. Transactions of the Wisconsin Academy of Sciences, Arts, and Letters
20:553 – 662.
Hertel, E. W. 1942. Studies on vigor in the rotifer Hydatina senta.
Physiological Zoology 15:304 – 324.
Herzig, A. 1983. Comparative studies on the relationship between
temperature and the duration of embryonic development of rotifers. Hydrobiologia 104:237 – 246.
Herzig, A. 1987. The analysis of planktonic rotifer populations: a
plea for long – term investigations. Hydrobiologia 147:163 – 180.
Hewitt, D. P., George, D. G. 1987. The population dynamics of Keratella cochlearis in a hypereutrophic tarn and the possible impact
of predation by young roach. Hydrobiologia 147:221 – 227.
8. Phylum Rotifera
Hillbricht – Ilkowska, A. 1983. Response of planktonic rotifers to the
eutrophication process and to the autumnal shift of blooms in
Lake Biwa, Japan. I. Changes in abundances and composition
of rotifers. The Japanese Journal of Limnology 44:93 – 106.
Hirata, H., Nagata, W. 1982. Excretion rates and excreted compounds of the rotifer Brachionus plicatilis O.F. Müller in culture. Memoirs of the Faculty of Fisheries Kagoshima University
31:161 – 174.
Hirayama, K., and S. Ogawa. 1972. Fundamental studies on the
physiology of the rotifer for its mass culture I. Filter feeding of
the rotifer. Bulletin of the Japanese Society of Scientific Fisheries
38:1207 – 1214.
Hofmann, W. 1977. The influence of abiotic factors on population
dynamics in planktonic rotifers. Archiv für Hydrobiologie Beiheft 8:77 – 83.
Hofmann, W. 1982. On the coexistence of two pelagic Filinia species
(Rotatoria) in Lake Plußee I. Dynamics of abundance and dispersion. Archiv für Hydrobiologie 95:125 – 137).
Hudson, C. T., Gosse, P. H. 1886. The Rotifera; or wheel-animalcules, both British and foreign. Vol. I & II. Longmans, Green,
London.
Hutchinson, G. E. 1967. A treatise on limnology. Vol. 2. Introduction to lake biology and the limnoplankton. Wiley, New York.
Hyman, L. H. 1951. The Invertebrates: Acanthocephala, Aschelminthes, and Entoprocta. The pseudocoelomate Bilateria.
Vol. 3. McGraw – Hill, New York.
Iltis, A., Riou – Duvat, S. 1971. Variations saisonniéres du peuplement en rotifères des eaux natronées du Kanem (Tchad). Cah.
O.S.T.R.O.M. ser. Hydrobiol. 5(2):101 – 112.
Ito, T. 1960. On the culture of the mixohaline rotifer Brachionus
plicatilis O.F. Müller, in seawater. Report of the Faculty of Fisheries, Prefectural University of Mie 3:708 – 740.
Jenkins, D. G., Buikema, A. L. 1985. Plankton rotifer responses to
herbicide stress in in situ microcosms. Virginia Journal of Science 36:144.
Jenkins, D.G., Underwood, M. O. 1998. Zooplankton may not disperse readily in wind, rain, or waterfowl. Hydrobiologia 387/
388:15 – 2.
Jennings, H. S., Lynch, R. S. 1928. Age, mortality, fertility and individual diversities in the rotifer Proales sordida Gosse. I. Effects
of the age of the parent on characteristics of the offspring. Journal of Experimental Zoology. 50:345 – 407.
Johansson, S. 1983. Annual dynamics and production of rotifers in
an eutrophication gradient in the Baltic Sea. Hydrobiologia
104:335 – 340.
Keshmirian, J., Nogrady, T. 1988. Histofluorescent labelling of catecholaminergic structures in rotifers (Aschelminthes) II. Males of
Brachionus plicatilis and structures from sectioned females. Histochemistry 89:189 – 192.
King, C. 1967. Food, age and the dynamics of a laboratory population of rotifers. Ecology 48:111 – 128.
King, C. E. 1969. Experimental studies of aging in rotifers. Experimental Gerontology 4:63 – 79.
King, C. E. 1972. Adaptation of rotifers to seasonal variation. Ecology 53:408 – 418.
King, C. E. 1977. Genetics of reproduction, variation, and adaptation in rotifers. Archiv für Hydrobiologie Beiheft 8:187 – 201.
King, C. E. 1980. The genetic structure of zooplankton populations.
in: Kerfoot, W. C. Ed., Evolution and ecology of zooplankton
communities. Univ. Press of New England, Hanover, NH, pp.
315 – 328.
King, C. E. 1983. A re – examination of the Lansing effect. Hydrobiologia 104:135 – 139.
King, C. E., Miracle, M. R. 1980. A perspective on aging in rotifers.
Hydrobiologia 73:13 – 19.
251
King, C. E., Snell, T. W. 1977. Culture media (natural and synthetic):
Rotifera in: M. Rechcigal, Jr., Ed. CRC Handbook Series in Nutrition and Food. Sect. G: Diets, Culture Media, Food Supplements. Vol II: Food habits of, and diets for invertebrates and
vertebrates — zoo diets. CRC Press, Cleveland, Oh, pp. 71 – 75.
King, C. E., Zhao, Y. 1987. Coexistence of rotifer (Brachionus plicatilis) clones in Soda Lake, Nevada. Hydrobiologia 147:57 – 64.
Kirk, K.L. 1991. Inorganic particles alter competition in grazing
plankton: the role of selective feeding. Ecology 72:915 – 923.
Kirk, K.L., Gilbert, J. J. 1992. Variation in herbivore response to
chemical defenses: zooplankton foraging on toxic cyanobacteria. Ecology 73:2208 – 2217.
Kirk, K. L. 1997. Life-history responses to variable environments:
starvation and reproduction in planktonic rotifers. Ecology 78:
434 – 441.
Kleinow, W. 1998. Stereopictures of internal structures and rotifer
trophi. Hydrobiologia 123 – 129.
Koste, W. 1976. Über die Rädertierbestände (Rotatoria) der oberen
und mittleren Hase in de Jahren 1966 – 1969. Osnabrücker
Naturwissenschaftliche. Mitteilungen 4:191 – 263.
Koste, W. 1978. Rotatoria. Die Rädertiere Mitteleuropas. 2 vols,
Gébrüder Borntraeger, Berlin.
Koste, W. 1989. Octotrocha speciosa, eine seltene, sessile Art mit
einem merkwürdigen Räderorgan. Mikrokosomos 78:115 – 121.
Koste, W., Shiel, R. J. 1986. Rotifera from Australian inland waters.
I. Bdelloidea (Rotifera: Digononta). Australian Journal of Marine and Freshwater Research 37:765 – 792.
Koste, W., Shiel, R. J. 1987. Rotifera from Australian inland waters.
II. Epiphanidae and Brachionidae (Rotifera: Monogononta). Invertebrate Taxonomy 7:949 – 1021.
Lorenzen, S. 1985. Phylogenetic aspects of pseudocoelomate evolution, in: Conway Morris, S., George, J. D., Platt, H. M. Eds.,
The origins and relationships of lower invertebrates. Systematics
Association, Vol. 28. Clarendon Press, Oxford, pp. 210 – 223.
Lubzens, E. 1987. Raising rotifers for use in aquaculture. Hydrobiologia 147:245 – 255.
Lubzens, E., Minkoff, G., Maron, S. 1985. Salinity dependence of
sexual and asexual reproduction in the rotifer Brachionus
plicatilis. Marine Biology 85:123 – 126.
Lubzens, E., Tandler, A., Minkoff, G. 1989. Rotifers as food in aquaculture. Hydrobiologia 186/187:387 – 400.
Luciani, A., Chasse, J. – L., Clément, P. 1983. Aging in Brachionus
plicatilis: the evolution of swimming as a function of age at two
different calcium concentrations. Hydrobiologia 104:141 – 146.
MacIsaac, H. J., Hutchinson, T. C., Keller, W. 1987. Analysis of
planktonic rotifer assemblages from Sudbury, Ontario, area
lakes of varying chemical composition. Canadian Journal of
Fisheries and Aquatic Sciences 44:1692 – 1701.
Magnien, R. E., Gilbert, J. J. 1983. Diel cycles of reproduction and
vertical migration in the rotifer Keratella crassa and their influence on the estimation of population dynamics. Limnology and
Oceanography 28:957 – 969.
Makarewicz, J. C., Likens, G. E. 1975. Niche analysis of a zooplankton community. Science 190:1000 – 1003.
Makarewicz, J. C., Likens, G. E. 1979. Structure and function of
the zooplankton community of Mirror Lake, New Hampshire.
Ecological Monographs 49:109 – 127.
Marcus, N. H., Lutz, R., Burnett, W., Cable, P. 1994. Age, viability,
and vertical distribution of zooplankton resting eggs from an
anoxic basin: Evidence of an egg bank. Limnology and
Oceanography 39:154 – 158.
Mark Welch, D. (2000). Evidence from a protein-coding gene that
acanthocephalans are rotifers. Invertebrate Biology:17–26.
May, L. 1980. On the ecology of Notholca squamula Müller in Loch
Leven, Kinross, Scotland. Hydrobiologia 73:177 – 180.
252
Robert Lee Wallace and Terry W. Snell
May, L. 1985. The use of procaine hydrochloride in the preparation
of rotifer samples for counting. Internationale Vereinigung für
theoretische und angewandte Limnologie, Verhandlungen 22:
2987 – 2990.
May, L. 1986. Rotifer sampling — a complete species list from one
visit? Hydrobiologia 134:117 – 120.
May, L. 1987. Effect of incubation temperature on the hatching of
rotifer resting eggs collected from sediments. Hydrobiologia
147:335 – 338.
May, L. 1989. Epizoic and parasitic rotifers. Hydrobiologia 186/187:
59 – 67.
Meadow, N. D., Barrows, Jr., C. H. 1971. Studies on aging in a bdelloid rotifer. Journal of Experimental Zoology 176:303 – 314.
Melone, G. 1998. The rotifer corona by SEM. Hydrobiologia 387/
388:131 – 134.
Melone, G., Ricci, C., Segers, H. 1998a. The trophi of Bdelloidea
(Rotifera): a comparative study across the class. Canadian Journal of Zoology 76:1755 – 1765.
Melone, G., Ricci, C., Segers, H., Wallace, R. L. 1998b. Phylogenetic
relationships of phylum Rotifera. Hydrobiologia 387/388:
101 – 107.
Miracle, M. R. 1974. Niche structure in freshwater zooplankton: a
principal components approach. Ecology 55:1306 – 1316.
Miracle, M. R., Serra, M., Vincente, E., Blanco, C. 1987. Distribution of Brachionus species in Spanish mediterranean wetlands.
Hydrobiologia 147:75 – 81.
Moore, M. V. 1988. Density-dependent predation of early instar
Chaoborus feeding on multispecies prey assemblages. Limnology and Oceanography 33:256 – 269.
Moore, M. V., Gilbert, J. J. 1987. Age-specific Chaoborus predation
on rotifer prey. Freshwater Biology 17:223 – 236.
Myers, F. J. 1937. A method of mounting rotifer jaws for study. Transactions of the American Microscopical Society 56:256 – 257.
Neill, W. E. 1984. Regulation of rotifer densities by crustacean zooplankton in an oligotrophic montane lake in British Columbia.
Oecologia 61:175 – 181.
Nipkow, F. 1961. Die Radertiere im Plankton des Zürchsee und
ihre Entwisklungsphasen. Schweizer Journal Hydrobiologie 23:
398 – 461.
Nogrady, T. 1982. Rotifera, in: Parker, S. P. Ed. Synopsis and classification of living organisms. McGraw – Hill., New York, pp.
865 – 872.
Nogrady, T., Alai. M.1983. Cholinergic neurotransmission in rotifers.
Hydrobiologia 104:149 – 153.
Nogrady, T., Keshmirian, J. 1986a. Rotifer neuropharmacology I.
Cholinergic drug effects on oviposition of Philodina acuticornis
(Rotifera, Aschelminthes). Comparative Biochemistry and Physiology 83C:335 – 338.
Nogrady, T., Keshmirian, J. 1986b. Rotifer neuropharmacology II.
Synergistic effect of acetyl – choline on local anesthetic activity
in Brachionus calyciflorus (Rotifera, Aschelminthes). Comparative Biochemistry and Physiology 83C:339 – 344.
Nogrady, T., Pourriot, R. 1995. Rotifera, Vol. 3: The Notommatidae.
in: Dumont, H. J., Ed., Guide to the identification of the microinvertebrates of the continental waters of the world. SPB
Academic Publishers, The Hague, The Netherlands, 248 pp.
Nogrady, T., Wallace, R. L., Snell, T. W. 1993. Rotifera: Vol. 1: Biology, Ecology and Systematics. in: Dumont, H. J. Ed., Guide to
the identification of the microinvertebrates of the continental
waters of the world. SPB Academic Publishers bv, The Hague,
The Netherlands, 142 pp.
O’Brien, W. J. 1987. Planktivory by freshwater fish: thrust and parry
in pelagia, in: Kerfoot, C. W., Sih, A. Eds. Predation: direct and
indirect impacts on aquatic communities. Univ. Press of New
England, Hanover, NH, pp. 3 – 16.
Pagani, M., Ricci, C., Bolzern, A. M. 1991. Comparison of five
strains of a parthenogenetic species, Macrotrachela quadricornifera (Rotifera, Bdelloidea) II. Isoenzymatice patterns. Hydrobiologia 211:157 – 163.
Paloheimo, J. E. 1974. Calculation of instantaneous birth rates. Limnology and Oceanography 19:692 – 694.
Pejler, B. 1956. Introgression in planktonic Rotatoria with some
points of view on its causes and results. Evolution 10:246 – 261.
Pejler, B. 1977a. General problems on rotifers taxonomy and global
distribution. Archiv für Hydrobiologie Beiheft 8:212 – 220.
Pejler, B. 1977b. On the global distribution of the family Brachionidae (Rotatoria). Archiv für Hydrobiologie (Suppl.) 53:
255 – 306.
Pennak, R. W. 1962. Quantitative zooplankton sampling in littoral
vegetation areas. Limnology and Oceanography 7:487 – 489.
Pennak, R. W. 1989. Freshwater invertebrates of the United States,
3rd ed. Wiley, New York.
Persoone, G., Van De Vel, A. Van Steertegem, M., De Nayer, B. 1989.
Predictive value of laboratory tests with aquatic invertebrates:
influence of experimental conditions. Aquatic Toxicology 14:
149 – 66.
Pilarska, J. 1972. The dynamics of growth of experimental populations of the rotifer Brachionus rubens Ehrbg. Polish Archives
for Hydrobiology 19:265 – 277.
Pontin, R. S. 1978. A key to the British freshwater planktonic Rotifera. Scientific Publications No. 38. Freshwater Biological Association, Cumbria, UK.
Pourriot, R. 1958. Sur l’ élevage des Rotifères au laboratoire. Hydrobiologia 11:189 – 197.
Pourriot, R. 1965. Recherches sur l’ecologie des Rotifères. Vie et Milieu (Suppl.) 21:1 – 224.
Pourriot, R. 1977. Food and feeding habits of Rotifera. Archiv für
Hydrobiologie Beiheft. 8:243 – 260.
Pourriot, R. 1979. Rotifères du sol. Revue d’Ecologie et de Biologie
du Sol. 16:279 – 312.
Pourriot, R. 1986. Les rotifères — Biologie. in: G. Barnabé, Ed. Aquaculture Vol. 1. Technique et documentation, Lavoisier, Paris.
Pourriot, R. 1997. Rotifera, Vol. 5: The Ituridae (Monogononta). in:
Dumont, H. J. Ed., Guides to the identification of the microinvertebrates of the continental waters of the world. SPB Academic Publishers, The Hague, The Netherlands. 344 pp.
Pourriot, R., Clément, P. 1981. Action de facteurs externes sur la reproduction et le cycle reproducteur des Rotifers. Acta Oecologia
Generale 2:135 – 151.
Pourriot, R., Rougier, C. 1975. Dynamique d’une population experimentale de Brachionus dimidatus (Bryce) (Rotifère) en fonction
de la nourriture et de la temperature. Annals Limnology 11:
125 – 143.
Pourriot, R., Rougier, C., Benest, D. 1986. Food quality and mictic
female control in the rotifer Brachionus rubens Ehr. Bulletin Society Zoologie Frances 111:105 – 112.
Preissler, K. 1980. Field experiments on the optical orientation of
pelagic rotifers. Hydrobiologia 73:199 – 203.
Rao, T. R., Sarma, S .S. S. 1986. Demographic parameters of Brachionus patulus Muller (Rotifera) exposed to sublethal DDT
concentrations at low and high food levels. Hydrobiologia 139:
193 – 200.
Ricci, C. 1983. Life histories of some species of Rotifera Bdelloida.
Hydrobiologia 104:175 – 180.
Ricci, C. 1984. Culturing of some bdelloid rotifers. Hydrobiologia
112:45 – 51.
Ricci, C. 1987. Ecology of bdelloids: how to be successful. Hydrobiologia 147:117 – 127.
Ricci, C. 1998a. Are lemnisci and proboscis present in the Bdelloidea?
Hydrobiologia 387/388:93 – 96.
8. Phylum Rotifera
Ricci, C. 1998b. Anhydrobiotic capabilities of bdelloid rotifers. Hydrobiologia 387/388:321 – 326.
Ricci, C., Vaghi, L., Manzini, M. L. 1987. Desiccation of rotifers
(Macrotrachela quadricornifera): survival and reproduction.
Ecology 68:1488 – 1494.
Ricci, C., Melone, G., Sotgia, C. 1993. Old and new data on
Seisonidea (Rotifera). Hydrobiologia 255/256:495 – 511.
Rico-Martinez, R., Snell, T. W. 1998. Mating behavior in eight rotifer species: using cross-mating tests to study species boundaries. Hydrobiologia 356:165 – 173.
Robb, E. J., Barron, G. L. 1982. Nature’s ballistic missile. Science
218:1221 – 1222.
Rothhaupt, K. O. 1985. A model approach to the population dynamics of the rotifer Brachionus rubens in two – stage chemostat culture. Oecologia 65:252 – 259.
Rougier, C., Pourriot, R. 1977. Aging and control of the reproduction in Brachionus calyciflorus (Pallas) (Rotatoria). Experimental Gerontology 12:137 – 151.
Ruttner – Kolisko, A. 1963. The interrelationships of the Rotatoria,
in: E. C. Dougherty, Ed., The Lower Metazoa. Univ. of California Press, Berkeley., CA, pp. 263 – 272.
Ruttner – Kolisko, A. 1969. Kreuzungexperimente zwischen Brachionus urceolaris and Brachionus quadridentatus, ein Beitrag
zur Fortpflanzungbiologie der heterogonen Rotatoria. Archiv
für Hydrobiologie 65:397 – 412.
Ruttner – Kolisko, A. 1974. Planktonic rotifers biology and taxonomy. Die Binnengewässer (Supplement) 26:1 – 146.
Ruttner – Kolisko, A. 1977a. The effect of the microsporid
Plistophora asperospora on Conochilus unicornis in Lunzer Untersee (LUS). Archiv für Hydrobiologie Beiheft 8:135 – 137.
Ruttner – Kolisko, A. 1977b. Amphoteric reproduction in a population of Asplanchna priodonta. Archiv für Hydrobiologie Beiheft
8:178 – 181.
Salt, G. W. 1987. The components of feeding behavior in rotifers.
Hydrobiologia 147:271 – 281.
Salt, G. W., Sabbadini, G. F., Commins, M. L. 1978. Trophi morphology relative to food habits in six species of rotifers (Asplanchnidae). Transactions of the American Microscopical Society 97:469 – 485.
Saunders, J. F. III, Lewis, Jr, W. M. 1989. Zooplankton abundance in
the lower Orinoco River, Venezuela. Limnol. Oceanogr. 34:
397 – 409.
Sawada, M., Enesco, H. E. 1984. A study of dietary restriction and
lifespan in the rotifer Asplanchna brightwelli monitored by
chronic neutral red exposure. Experimental Gerontology 19:3
29 – 334.
Schlüter, M. 1980. Mass culture experiments with Brachionus
rubens. Hydrobiologia 73:45 – 50.
Schlüter, M., Groeneweg, J. 1981. Mass production of freshwater rotifers on liquid wastes. I. The influence of some environmental
factors on population growth on Brachionus rubens Ehrenberg
1838. Aquaculture 25:17 – 24.
Schlüter, M., Groeneweg, J. 1985. The inhibition by ammonia of
population growth of the rotifer, Brachionus rubens, in continuous culture. Aquaculture 46:215 – 220.
Schlüter, M., Groeneweg, J., Soeder, C. J. 1987. Impact of rotifer grazing on population dynamics of green microalgae in high – rate
ponds. Water Research 10:1293 – 1297.
Schmid-Araya, J. M. 1998. Rotifers in interstitial sediments. Hydrobiologia 231 – 240.
Schnese, W. 1973. Relations between phytoplankton and zooplankton in brackish coastal water. Oikos (Suppl.) 15:28 – 33.
Schramm, U., Becker, W. 1987. Anhydrobiosis of the bdelloid
rotifer Habrotrocha rosa (Aschelminthes). Zeitschrift fuer
Mikroskopisch – Anatomische Forschung 101:1 – 17.
253
Scott, J. M. 1983. Rotifer nutrition using supplemented monoxenic
cultures. Hydrobiologia 104:155 – 166.
Scott, J. M. 1988. Effect of growth rate on the physiological rates of
a chemostat – grown rotifer Encentrum linnhei. Journal of the
Marine Biological Association of the United Kingdom 68:
165 – 177.
Seaman, M. T., Gophen, M., Cavari, B. Z., Azoulay, B. 1986. Brachionus calyciflorus Pallas as agent for removal of E. coli in
sewage ponds. Hydrobiologia 135:55 – 60.
Segers, H. 1995a. Rotifera, Volume 2: The Lecanidae (Monogononta).,
in: Dumont, H. J., Ed. Guides to the identification of the microinvertebrates of the continental waters of the world. SPB Academic
Publishers, The Hague, The Netherlands. 226 pp.
Segers, H. 1995b. Rotifera, Volume 3: The Scaridiidae. in: Dumont,
H. J., Ed., Guides to the identification of the microinvertebrates
of the continental waters of the world. SPB Academic Publishers, The Hague, The Netherlands. 248 pp.
Serra, M., Miracle, M. 1985. Enzyme polymorphisms in Brachionus
plicatilis populations from several Spanish lagoons. Internationale Vereinigung für theoretische und angewandte Limnologie, Verhandlungen 22:2991 – 2996.
Serrano, L., Miracle, M. R., Serra, M. 1986. Differential response of
Brachionus plicatilis (Rotifera) ecotypes to various insecticides.
Journal of Environmental Biology 7:259 – 275.
Sladecek, V. 1983. Rotifers as indicators of water quality. Hydrobiologia 100:169 – 201.
Snell, T. W. 1977. Lifespan of male rotifers. Archiv für Hydrobiologie Beiheft 8:65 – 66.
Snell, T. W. 1979. Intraspecific competition and population structure
in rotifers. Ecology 60:494 – 502.
Snell, T. W. 1980. Blue – green algae and selection in rotifer populations. Oecologia 46:343 – 346.
Snell, T. W. 1986. Effects of temperature, salinity and food level on
sexual and asexual reproduction in Brachionus plicatilis (Rotifera). Marine Biology 92:157 – 162.
Snell, T. W. 1989. Systematics, reproductive isolation and species
boundaries in monogonont rotifers. Hydrobiologia 186/187:
299 – 310.
Snell, T. W., Childress, M. J. 1987. Aging and loss of fertility in male
and female Brachionus plicatilis (Rotifera). International Journal
of Invertebrate Reproduction and Development 12:103 – 110.
Snell, T. W., Garman, B. L. 1986. Encounter probabilities between
male and female rotifers. Journal of Experimental Marine Biology and Ecology 97:221 – 230.
Snell, T. W., Hawkinson, C. A. 1983. Behavioral reproductive isolation among populations of the rotifer Brachionus plicatilis. Evolution 37:1294 – 1305.
Snell, T. W., Hoff, F. H. 1985. The effect of environmental factors on
resting egg production in the rotifer Brachionus plicatilis. Journal of the World Mariculture 16:484 – 497.
Snell, T. W., Hoff, F. H. 1987. Fertilization and male fertility in the
rotifer Brachionus plicatilis. Hydrobiologia 147:329 – 334.
Snell, T. W., King, C. E. 1977. Lifespan and fecundity patterns in rotifers: the cost of reproduction. Evolution 31:882 – 890.
Snell, T. W., Winkler, B. C. 1984. Isozyme analysis of rotifer protein.
Biochemical Systematics and Ecology 12:199 – 202.
Snell, T. W., Burke, B. E., Messur, S. D. 1983. Size and distribution of
resting eggs in a natural population of the rotifer Brachionus
plicatilis. Gulf Research Reports 7:285 – 288.
Snell, T. W., Childress, M. J., Boyer, E. M., Hoff, F. H. 1987. Assessing the status of rotifer mass cultures. Journal of the World
Aquaculture Society 18:270 – 277.
Snell, T. W., Childress, M. J., Winkler, B. C. 1988. Characteristics of
the mate recognition factor in the rotifer Brachionus plicatilis.
Comparative Biochemistry and Physiology 89A:481-485.
254
Robert Lee Wallace and Terry W. Snell
Snell, T. W., Janssen, C. R. 1995. Rotifers in ecotoxicology: a review.
Hydrobiologia 313/314:231 – 247.
Snell, T. W., Janssen, C. R. 1998. Microscale toxicity testing with rotifers, in: Wells, P. G., Lee, K., Blaise, C. Eds., Microscale Toxicity Testing in Aquatic Toxicology, CRC Press, Boca Raton, FL,
pp. 409 – 422.
Snell, T. W., Rico-Martinez, R., Kelly, L., N. Battle, T. E. 1995. Identification of a sex pheromone from a rotifer. Marine Biology
123: 347 – 353.
Starkweather, P. L. 1980. Aspects of the feeding behavior and trophic
ecology of suspension feeding rotifers. Hydrobiologia 73:63 – 72.
Starkweather, P. L. 1987. Rotifera. Pages 159 – 183 in: T. J. Pandian
and F. J. Vernberg, Eds., Animal energetics. Vol. 1: Protozoa
through Insecta. Academic Press, Orlando, FlO.
Starkweather, P. L., Kellar, P. E. 1987. Combined influences of particulate and dissolved factors in the toxicity of Microcyctis aeruginosa (NRC – SS – 17) to the rotifer Brachionus calyciflorus. Hydrobiologia 147:375 – 378.
Stemberger, R. S. 1979. A guide to rotifers of the Laurentian Great
Lakes. U.S. Environmental Protection Agency, Cincinnati, Oh.
(Available from National Technical Information Service, Springfield, VA, PB80 – 101280.)
Stemberger, R. S. 1981. A general approach to the culture of planktonic rotifers. Canadian Journal of Fisheries and Aquatic Sciences 38:721 – 724.
Stemberger, R. S. 1984. Spine development in the rotifer Keratella
cochlearis: induction by cyclopoid copepods and Asplanchna.
Freshwater Biology 14:639 – 647.
Stemberger, R. S. 1988. Reproductive costs and hydrodynamic benefits of chemically induced defenses in Keratella testudo. Limnology and Oceanography 33:593 – 606.
Stemberger, R. S. 1990. Food limitation, spination, and reproduction
in Brachionus calyciflorus. Limnology and Oceanography 35:
33 – 44.
Stemberger, R. S., Gilbert, J. J. 1984a. Body size, ration level, and population growth in Asplanchna. Oecologia (Berlin) 64:355 – 359.
Stemberger, R. S., Gilbert, J. J. 1984b. Spine development in the rotifer Keratella cochlearis: induction by cyclopoid copepods and
Asplanchna. Freshwater Biology 14:639 – 647.
Stemberger, R. S., Gilbert, J. J. 1985. Body size, food concentration and
population growth in planktonic rotifers. Ecology 66:1151 – 1159.
Stemberger, R. S., Gilbert, J. J. 1987a. Defenses of planktonic rotifers
against predators. in: Kerfoot, W. C., Sih, A. Eds. Predation: direct and indirect impacts on aquatic communities. Univ. Press of
New England, Hanover, NH, pp. 227 – 239.
Stemberger, R. S., Gilbert, J. J. 1987b. Multiple species induction of
morphological defenses in the rotifer Keratella testudo. Ecology
68:370 – 378.
Taylor, H. L. 1993. The Taylor microcompressor Mark II. Microscope 41:19-20.
Threlkeld, S. T., Rybock, J. T., Morgan, M. D., Folt, C. L., Goldman,
C. R. 1980. The effects of an introduced invertebrate predator
and food resource variation on zooplankton dynamics in an ultraoligotrophic lake, in: Kerfoot, W. C. Ed., Evolution and Ecology of Zooplankton Communities. Univ. Press of New England,
Hanover, NH, pp. 555 – 568.
Tzean, S. S., Barron, G. L. 1983. A new predatory hypomycete capturing bdelloid rotifers in soil. Canadian Journal of Botany
61:1345 – 1348.
Wallace, R. L. 1977. Distribution of sessile rotifers in an acid bog
pond. Archiv für Hydrobiologie 79:478 – 505.
Wallace, R. L. 1980. Ecology of sessile rotifers. Hydrobiologia 73:
181 – 193.
Wallace, R. L. 1987. Coloniality in the phylum Rotifera. Hydrobiologia 147:141 – 155.
Wallace, R. L. 1998. Rotifera, in: Knobil, E., Neil. J. D., Eds., Encyclopedia of reproduction. Vol. 4. Academic Press, San Diego,
CA, pp. 118 – 129.
Wallace, R. L., Edmondson, W. T. 1986. Mechanism and adaptive
significance of substrate selection by a sessile rotifer. Ecology
67:314 – 323.
Wallace, R. L., Taylor, W. K. Litton, J. R. 1989. Invertebrate zoology.
Macmillan, New York. 337 pp.
Wallace, R. L., Ricci, C., Melone, G. 1996. A cladistic analysis of
pseudocoelomate (aschelminth) morphology. Invertebrate Biology 115:104 – 112.
Walsh, E. J. Zhang, L. 1992. Polyploidy and body size variation in a
natural population of the rotifer Euchlanis dilatata. Journal
Evolutionary Biology 5:345 – 353.
Walz, N. 1983a. Continuous culture of the pelagic rotifers Keratella
cochlearis and Brachionus angularis. Archiv für Hydrobiologie
98:70 – 92.
Walz, N. 1983b. Individual culture and experimental population dynamics of Keratella cochlearis (Rotatoria). Hydrobiologia 107:
35 – 45.
Walz, N. 1987. Comparative population dynamics of the rotifers
Brachionus angularis and Keratella cochlearis. Hydrobiologie
147:209 – 213.
Walz, N. (Ed.) 1993. Plankton regulation dynamics. Springer, New
York, 308 pp.
Walz, N. 1995. Rotifer populations in plankton communities: energetics and life history strategies. Experentia 51:437 – 453.
Walz, N. 1997. Rotifer life history strategies and evolution in freshwater plankton communities, in: Streit, B., Städler, T., Lively, C.M.
Eds., Evolutionary Ecology of Freshwater Animals. Birkhäuser
Verlag, Basel, pp. 119 – 149.
Walz, N., Elster, H. – J. Mezger, M. 1987. The development of the rotifer community structure in Lake Constance during its eutrophication. Archiv für Hydrobiologie (Suppl) 74:452 – 487.
Watanabe, T., Kitajima, C., Fujita, S. 1983. Nutritional values of live
food organisms used in Japan for mass propagation of fish: a review. Aquaculture 34:115 – 143.
Williamson, C. E. 1983. Invertebrate predation on planktonic rotifers.
Hydrobiologia 104:385 – 396.
Williamson, C. E. 1987. Predator – prey interactions between omnivorous diaptomid copepods and rotifers: the role of prey morphology and behavior. Limnology and Oceanography 32:167 – 177.
Wurdak, E. 1987. Ultrastructure and histochemistry of the stomach
of Asplanchna sieboldi. Hydrobiologia 147:361 – 371.
Wurdak, E., Clément, P., Amselem, J. 1983. Sensory receptors involved
in the feeding behavior of the rotifer Asplanchna brightwelli.
Hydrobiologia 104:203 – 212.
Wurdak, E., Gilbert, J. J. 1976. Polymorphism in the rotifer Asplanchna sieboldi: fine structure of saccate, cruciform and campanulate females. Cell Tissue Research 169:435 – 448.
Wurdak, E., Gilbert, J. J., Jagels, R. 1977. Resting egg ultrastructure
and formation of the shell in Asplanchna sieboldi and Brachionus
calyciflorus. Archiv für Hydrobiologie Beiheft 8:298 – 302.
9
NEMATODA AND NEMATOMORPHA
George O. Poinar, Jr.
Department of Entomology
Oregon State University
Corvallis, Oregon 97331
NEMATODA
I. Introduction
II. Morphology and Physiology
A. Cuticle
B. Hypodermis
C. Muscular System
D. Digestive System
E. Excretory System
F. Respiration
G. Nervous System
H. Sense Organs
I. Reproductive System
III. Development and Life History
IV. Ecology
A. Fossil Record
B. Habitats of Freshwater Nematodes
C. Enemies
V. Collecting and Rearing Techniques
A. Sampling Methods
B. Extraction
NEMATODA
I. INTRODUCTION
The phylum Nematoda comprises a wide range of
roundworms that can be grouped either in nutritional
(microbotrophic, predaceous, and parasitic in plants,
invertebrates, or vertebrates) or ecological (soil, marine, freshwater, ectoparasitic, endoparasitic, or freeliving) categories (Poinar, 1983). Relatively little attention has been devoted to this important group of
invertebrates by freshwater biologists because of difficulties associated with sampling, extraction, and identification.
Nematode representatives of essentially all of
the nutritional categories mentioned above occur in
freshwater during one or more life stages. However
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
C. Fixing and Mounting
D. Culturing
VI. Identifcation
A. Taxonomic Key to Families
and Selected Genera of Freshwater
Nematoda
B. Taxonomic Key to Extant Genera
of Freshwater Mermithidae
NEMATOMORPHA
VII. Introduction
VIII. Morphology and Physiology
IX. Development and Life History
X. Sampling
XI. Identification
A. Taxonomic Key to Extant Genera
of Nematomorpha
Literature Cited
Appendix 9.1 Systematic Arrangement
of the Nematode Genera
nematodes are treated here only if all or a large part of
their life cycle occurs in freshwater habitats (these habitats are discussed elsewhere). Nematode parasites of
vertebrates that live in or frequent freshwater usually
occur in the freshwater habitat only as eggs or within
intermediate hosts and, therefore, are not discussed in
this chapter. Mermithid parasites of insects are included since the eggs, infective stages, postparasitic
juveniles, and adults occur in freshwater habitats.
Only nematode genera containing obligate aquatic
species (those which live nowhere else) that have been
reported from North America are included in the keys
in the present work. The systematic arrangement of
these genera is listed in Appendix 9.1. As more studies
are undertaken, it is likely that genera described from
other continents will also be represented in North
America. In addition, the reader should note that
255
256
George O. Poinar, Jr.
undescribed or unreported genera may be encountered
in routine sampling.
The related group, Nematomorpha, or hairworms,
are frequently encountered by freshwater biologists.
Superficially, they resemble mermithid nematodes, and
share similar life history strategies with certain species
of the latter group.
II. MORPHOLOGY AND PHYSIOLOGY
Nematodes are nonsegmented, wormlike invertebrates lacking jointed appendages but possessing a
body cavity and a complete alimentary tract (Fig. 1).
While they lack specialized respiratory and circulatory
systems, nematodes possess a well-developed nervous
system, an excretory system, and a set of longitudinal
muscles.
With the exception of the family Mermithidae,
most freshwater nematodes are under 1 cm in length.
Although the basic body shape and anatomic plan of
all nematodes are similar, there are some characteristics
of most freshwater nematodes that are lacking in many
terrestrial forms. One of these is the presence of three
unicellular hypodermal glands commonly located in the
upper part of the tail, immediately behind the anus
(Figs. 1 and 2D). These caudal glands produce a secretion that is carried by a canal to the tip of the tail. At
this point, the canal is attached to a valve (spinneret),
which passes through a pore in the tail terminus (Figs.
1 and 6d). The secretion produces an adhesive deposit
by which the nematodes can attach themselves to the
surfaces of submerged rocks, plants, invertebrates, and
debris.
Mucus may also be produced by pharyngeal glands
of aquatic nematodes. The deposits from the caudal
and pharyngeal glands can be used to form slimy traces
over the substrate. Microorganisms that become entrapped in these deposits serve as food when the nematodes ingest the mucus together with the attached
microorganisms (Riemann and Schrage, 1978).
A. Cuticle
The exterior body covering on all nematodes is
a noncellular, flexible, multilayered structure called
FIGURE 1 Adult female of Plectus (Plectidae) [center insert shows tail of male]. M, mouth; S, stoma; P, pharynx; N, nerve ring; E, excretory pore; B, valvated basal bulb of pharynx; I, intestine; R, rectum; A, anus; C,
caudal glands; T, spinneret; V, vulva; O, ovary; F, egg; D, spicule; G, gubernaculum; L, genital papillae. [Drawing by A. Maggenti, Univeristy of California, Davis; labeling by G. Poinar.]
FIGURE 2 (a) Gradually tapering pharynx of a freshwater Dorylaimus sp. Note absence of distinct
valve in basal portion of pharynx. Arrow shows the nerve ring. (b) Fossil tracts thought to be made
by freshwater nematodes. From the 50 million-year-old (Middle Eocene) Green River sediments of
Lake Uinta in the Soldier summit area near Provo, Utah. (c) A juvenile freshwater nematode capable
of swimming by rapid vibratory body movements. (d) Tail of a freshwater nematode showing three
caudal glands (CG), the caudal gland tube (T) and spinneret (arrow).
257
258
George O. Poinar, Jr.
the cuticle. It is secreted by a layer of underlying hypodermal cells and covers the entire external surface of
the nematode, with portions entering various external
openings such as the stoma, rectum, vagina, amphids
(lateral sense organs), and sometimes the excretory
pore. The cuticle is composed of an outer cortex, a
middle matrix, and an inner fiber or basal zone. All
these regions are constructed of sublayers, which vary
in thickness according to the species and stage. The
outermost lipoidal layer of the cortex is important
because it serves as a semipermeable membrane, allowing water and solutes to pass in and out of the
body cavity.
The cuticle of freshwater nematodes may bear longitudinal striations, punctations, bristles, setae, or somatic alae (Figs. 6b and 7a). When present, the bursa is
attached to the male tail and is composed of flattened
flaps of cuticle which guide and hold the male in place
during mating (Fig. 6c). All these cuticular structures
are useful characters for identifying nematode groups
or species. The cuticle is normally shed four times during development. At each molt (ecdysis), the old cuticle
splits and is discarded. In a few forms, the old cuticle is
retained, producing a double cuticle or sheath around
the nematode (Fig. 8b).
B. Hypodermis
The hypodermis is a layer of tissue that is located
beneath the cuticle and is responsible for the formation
of the cuticle. It is generally a thin layer of tissue that
expands into the coelom to form longitudinal cords
between the muscle fields. Most nematodes possess lateral, dorsal, and ventral cords that contain the nuclei
and other cytoplasmic inclusions of the hypodermis.
The hypodermal cords originate as distinct cells but
later lose their identity and form a syncytium.
C. Muscular System
Nematode movement is controlled by longitudinal
muscles because these organisms do not possess circular muscles. These somatic muscles are composed of
many adjacent cells which are innervated by nerves
lodged in the ventral and dorsal hypodermal cords.
Most nematode movement superficially resembles that
of many other elongate, stiff-bodied, appendageless
animals, whether they be vertebrates (snakes) or invertebrates (biting midge larvae). In locomotion, the sideto-side bending of the body is produced by alternating
contractions and relaxations of opposite muscle sectors
that are controlled by the central nervous system.
When placed on flat surfaces in the laboratory, nematodes normally crawl on their sides. Some aquatic
nematodes can swim by rapid vibratory or thrashing
side-to-side movements of the body.
D. Digestive System
All freshwater nematodes, except the Mermithidae,
contain a continuous alimentary tract composed of a
stoma, pharynx, and intestine (Fig. 1). The stoma
(mouth) is the most anterior part of the digestive tract
and is usually lined with cuticle. The stomal walls are
formed by a fusion of various smaller segments or rings
(rhabdions), which may be separated as is found in the
Cephalobidae, or fused, as in the Rhabditidae (Fig. 8a).
The structure of the stoma generally reflects the food
selection of the bearer. For example, predaceous nematodes tend to have a wide stoma armed with teeth
(Fig. 7b), a stylet, or a spear (Fig. 9b), whereas microbotrophic forms generally have a small tubular stoma
(Fig. 8a), and omnivores a funnel-shaped mouth cavity
(Fig. 10a). Plant parasites have protrusible stylets
which can be inserted into plant tissues (Fig. 8b), and
the infective stages of invertebrate parasites often have
a small stylet used in penetrating the cuticle or intestinal wall of the host.
The pharynx (or esophagus) lies between the stoma
and intestine and passes food from the mouth into the
intestine. It is usually composed of radial muscles,
which upon dilation allow the pharynx to function as a
pump or vacuum. The pharynx also contains glands
that produce secretions used in digestion, escape from
the egg shell, molting, and penetration of host tissue.
There are normally three portions to the pharynx: (1)
the corpus (sometimes enlarged to form a distinct
metacorpus or median bulb); (2) the isthmus (usually
surrounded by the nerve ring); and (3) the basal bulb
(often containing valves to keep food particles from reversing their direction of flow) (Fig. 8a). When nematodes feed, material is sucked up into the mouth by reduced pressure resulting from contraction of the radial
muscles of the pharynx. Food is passed along the lumen by waves of alternating contractions and relaxations of the pharyugeal muscles. The pharyngeal lumen opens behind the food, thus creating suction
pressure, and closes in front of the food, forcing it
through.
The intestine is a single-cell-thick, cylindrical tube
that runs from the pharynx to the anal opening (cloacal
opening in males). It is composed of an anterior ventricular region, a midintestine or intestine proper, and a
posterior rectal region (not always obvious). The inner
surfaces of the cells facing the lumen are lined with
microvilli which seem to be responsible for the major
uptake of nutrients. A pharyngeal-intestinal valve is
located in the anterior portion, behind the basal bulb
9. Nematoda and Nematomorpha
of the pharynx. In some freshwater forms, this valve is
quite distinct and elongated (10c). An intestinal – rectal
valve is located at the junction of the intestine and rectum. This unicellular sphincter muscle controls the
amount of water passing out of the body. Many rhabditid-type nematodes have three rectal glands, which
empty into the rectum. These glands are not to be confused with the caudal glands, which produce secretions
from the tail terminus (Fig. 2d).
The Mermithidae are unusual in having the intestine detached from the remainder of the alimentary
tract and modified into a trophosome or food storage
organ. During development, the trophosome separates
from the pharynx and rectum, thus remaining as a separate, isolated organ.
E. Excretory System
Although variable in structure, the excretory system of nematodes falls into two basic types. The first is
frequently found in freshwater forms and consists of a
ventral excretory gland (renette cell) connected by an
excretory duct to the excretory pore on the surface of
the cuticle. The ventral pore usually opens in the pharyngeal or anterior intestinal region of the body. The
second type is a tubular system consisting of a series of
longitudinal excretory canals; pooled contents of these
canals pass into a joining canal which connects with
the excretory duct and pore. Most freshwater nematodes excrete nitrogen in the form of ammonia or urea.
The excretory system also has an osmoregulatory function of removing water that diffuses into the pseudocoelom and so regulates the amount of turgor pressure
in the body cavity; some scientists believe that this is
the primary function of the “excretory” system.
F. Respiration
Most nematodes are aerobic organisms, at least
during their developmental period. Many nematodes
can survive short periods of anaerobic conditions but
only a few forms can survive anoxia indefinitely.
Strayer (1985) listed representatives of three nematode
genera that he considered as benthic species found under anoxic conditions in Mirror Lake. A specialized
respiratory system is lacking in nematodes. Instead,
they obtain and lose gases by simple diffusion. Body
cavity cells, when present, appear to play no role in
oxygen transport.
G. Nervous System
The nervous system of nematodes centers around a
central anterior mass of ganglia or “brain,” which
259
connects with nerve cords extending anteriorly and
posteriorly through the body. The central ganglionic
mass is closely associated with the circumpharyngeal
commissure or nerve ring (Fig. 2a). Nerves extending
anteriorly from the nerve ring innervate the amphids
and cephalic sense organs, while those running posteriorly control the male tail papillae.
H. Sense Organs
The size, shape, position, and spatial arrangement
of sense organs are all characteristic of nematode
species or groups and are therefore frequently used as
taxonomic characters. The anterior sensory papillae are
found on the nematode head. The basic number is 16,
but usually some have become vestigial. There is normally an inner circle of six labial papillae around the
mouth, a second ring of six labial papillae slightly behind the first set, and a third circle of four cephalic
papillae still further back on the head. In mermithids,
the cephalic papillae may also include adjacent enlargements of the hypodermal tissue.
The amphids are paired, lateral sense organs
that open to the exterior on the nematode cuticle.
They may be located on the same circle as the labial
or cephalic papillae or further back in the neck region. The term amphid generally refers to the exterior
configuration of the amphidial opening, although the
amphid also consists of an amphidial pouch, amphidial gland, and amphidial nerves. The structure of
the amphidial openings and their location are very
important taxonomic characters. They can vary from
small pores to large circles (Fig. 7c), pockets (Fig.
10b), or spirals (Fig. 7d). Sexual dimorphism may
be evident with either the male or female possessing
a considerably larger amphid usually of the same
general shape. Amphids may be multifunctional and
evidence suggests that they serve as photoreceptors,
olfactors, or sensory glands. Some are sensitive to pH
and ions.
Other innervated papillae on nematodes are deirids
(paired papillae located laterally near the nerve ring),
postdeirids (similar structures located at the midbody),
and genital papillae (found on the ventral surface
around the cloacal opening of males). When elongate,
the genital papillae are usually attached to thin membranous outgrowths of cuticle (bursa) (Fig. 6c). The
presence or absence of a bursa and its structure are important taxonomic characters.
A few freshwater nematodes possess what are
thought to be light receptor organs. These receptors
normally consist of pigmented areas in the neck region, which may or may not be associated with a
cuticular-structured lens. If a lensatic body is present,
260
George O. Poinar, Jr.
then the organ is sometimes referred to as an ocellus or
pseudocellus.
I. Reproductive System
The majority of nematodes, including the freshwater forms, reproduce by amphimixis (sperm and eggs
come from separate individuals). However, some forms
have developed uniparental reproduction (i.e., autotoky), and this condition also occurs in aquatic genera.
In the latter, autotoky usually expresses itself as
parthenogenesis, where progeny arise from unfertilized
eggs. Hermaphroditism seems quite rare among freshwater nematodes (Poinar and Hansen, 1983), apparently occurring in Chronogaster troglodytes (Poinar
and Sarbu, 1994) and some others. Asexual reproduction does not occur in the Nematoda.
In amphimictic reproduction, the male nematode
always places sperm inside the female; external fertilization has never been demonstrated. Some aquatic
species exhibit “traumatic insemination” when the
male penetrates the female cuticle with his spicules and
releases sperm into her body cavity.
Female nematodes contain one or two gonads,
which open to the exterior on the ventral side of the
body at the vulva (Figs. 1 and 6a). The vulva is connected to a muscular tube, the vagina, which in turn
leads into the uterus followed by the oviduct and
ovary. A spermatheca (sperm-collecting area) is usually
present between the uterus and oviduct. With doubleovary species (didelphic), the gonads are usually opposite and join at a common vagina (Fig. 6a). Singleovary forms (monodelphic) have retained the anterior
ovary, which is often reflexed into the posterior part of
the body (Fig. 8d).
The male may have a single testis (monorchic) or
two testes (diorchic), which lead to a common seminal
vesicle and vas deferens before entering the cloacal chamber, a common opening for the reproductive and digestive systems. Males of freshwater nematodes all possess
one or two sclerotized structures termed spicules (Figs. 1
and 6c). These are inserted into the vagina of the female
at insemination (except in traumatic insemination when
they penetrate the cuticle of the female). Some males also
have another sclerotized structure, the gubernaculum.
The spicules are contained within the walls of the cloacal
chamber while the gubernaculum is attached to the floor
of the cloacal chamber. The latter structure supports the
spicules during their movement in and out of the cloacal
opening (Fig. 6c). On the ventral surface of the male tail
is usually some type of genital papillae, which may or
may not be supported by a bursa (Fig. 6c).
Through the release and detection of sex attractants, male and female nematodes are able to locate
each other. Once together, the nematodes intertwine
and the posterior region of the male contacts the female vulva. Sometimes an adhesive compound is produced to seal the union. Spicules are then inserted and
sperm is transferred from the vas deferens of the male
into the vagina and uterus of the female. The pair may
remain joined for only several minutes or may be
united for days. It is common to find “balls” of
mermithids in stream bottoms all coiled together in
continuous copulation. Individual sperm as well as secondary spermatocytes may be transferred during mating. In many nematodes, sperm maturation is completed in the female. Nematode sperm is variable in size
and shape, but is always nonflagellated and usually
amoeboid.
Parthenogenetic forms can reproduce by mitotic or
meiotic parthenogenesis. In the former, the diploid somatic number of chromosomes is retained, whereas in
the latter, two maturation divisions occur which are
similar to oogenesis in amphimictic species. A diploid
chromosome number is established by fusion of the
nonextruded polar nucleus with the egg pronucleus or
by doubling of the chromosome before the first cleavage division occurs.
III. DEVELOPMENT AND LIFE HISTORY
The eggs of nematodes differ from all other stages
in containing chitin in their shells. During embryonic
development, nematodes show predeterminate cleavage, which means that cells destined to form specific
tissues appear early in the embryo. Freshwater forms
like the mermithids may deposit eggs with fully
formed juveniles ready to hatch upon receiving the
right stimuli. Other species deposit eggs in the singlecell stage, with embryonic development occurring in
the environment. Postembryonic development in nematodes (development after hatching) is similar to the
gradual type of metamorphosis in insects. Aside from
an increase in size, proportional changes of various organs, and the development of the gonads and second
sexual characteristics, there are few differences between juvenile and adult nematodes (Fig. 11c). For this
reason, the immature stages of nematodes should be
called juveniles and not larvae, because the latter usually implies complete metamorphosis where the immature stages differ radically from the adults. [Despite
these differences, however, the term larva is widely
used today in referring to the immature, postembryonic stages of nematodes.]
Hatching is influenced by temperature and external
stimuli. The eggs of the mermithid Pheromermis
pachysoma hatch when ingested by aquatic insects.
9. Nematoda and Nematomorpha
Eggs of other freshwater nematodes hatch when the
water reaches a certain temperature. Eggs that undergo
anabiosis as a result of desiccation will not hatch until
the area is flooded. This probably enhances survival for
nematodes in temporary water sources, although this
point still requires elucidation. In most cases, it is the
first-stage juvenile that emerges from the egg; however,
in mermithids, juveniles molt once in the egg and then
emerge as second-stage juveniles. If equipped with a
spear or tooth, the young nematode will use these to
break out of the shell, otherwise it simply employs
pressure and pharyngeal secretions to soften the shell
wall.
All nematodes undergo four molts and have six
stages during their development (i.e., egg, first-stage juvenile, second-stage juvenile, third-stage juvenile,
fourth-stage juvenile, and adult). During each molt, the
cuticle is shed and replaced by another secreted by the
hypodermis. Sometimes two molts may occur almost
simultaneously, as in postparasitic juvenile Mermithidae (Fig. 9a), and occasionally the shed skin is retained
around the body of the next stage (Fig. 8b). Free-living
nematodes generally complete their development
rapidly in comparison to other metazoans. This can be
3 – 5 days for rhabditids (under optimum conditions)
and generally from 1 to 6 weeks for most other freshwater forms. Therefore, nematode populations can
turn over very quickly (see Schiemer, 1983, 1987 for
energetic studies on nematodes).
Aside from premolt and quiescent stages, most nematodes feed continuously throughout the growth period. Most freshwater nematodes take food in through
the mouth and absorb nutrients into the intestinal cells.
During their parasitic development in the hemolymph
of invertebrates, mermithids absorb nutrients directly
through their body walls. Nematode growth entails
both enlargement and multiplication of cells. Growth is
frequently nonproportional from stage to stage. For
example, the intestine often will grow faster than the
pharynx, so the proportion of the two organs differs in
each stage. This information can be used to determine
nematode stages, along with gonad development.
The types of food vary among the freshwater nematode groups. Microbotrophic forms ingest bacteria,
algae, single-celled fungi, and protozoa. Predaceous
species attack small metazoans such as nematodes,
annelids, early insect stages, and molluscs. Many are
omnivorous and will consume microbial life as well as
metazoans. Plant parasites feed on the cytoplasm of
higher plants, algae and filamentous fungi. Parasites
of invertebrates absorb nutrients from the hemolymph
of various taxa, especially insects in the case of the
Mermithidae and snails in the case of Daubaylia. Vertebrate parasites may occur in the guts or body cavities
261
of fish, aquatic birds, or amphibians, where they feed
on host fluids and tissues. Only the egg stages are freeliving and these are not likely to be encountered or recognized during routine collecting activities.
The dependence of nematodes on moisture has
brought a tremendous selection pressure to withstand
periods of desiccation. Some nematodes form resistant
stages, while in others, all life stages can tolerate
drought conditions for various periods. Resistant
stages include the egg and the second- through fourthstage juveniles. Some nematodes have been maintained
for years (up to 25 years) in anabiosis and then revived
in water.
Very little is known about the resistant stages, dispersal, and survival of freshwater nematodes. Jacobs
(1984) mentions the “ability for passive dispersal” of
resistant forms, but does not discuss how this dispersal
occurs. It is very likely that the eggs and, possibly,
some juvenile stages of forms adapted to semitemporary water sources can enter a resistant, quiescent stage
capable of surviving desiccation and possibly temperature extremes; these have been demonstrated in some
soil and plant nematodes. Passive dispersal may occur
when these resistant stages are blown by heavy winds
or washed to new areas by flash floods. The possibility
of freshwater nematodes being carried in mud attached
to the body parts of various water-frequenting animals,
such as wading and swimming birds, is also feasible.
The presence of freshwater nematodes in temporary
ponds has not been investigated.
IV. ECOLOGY
A. Fossil Record
The oldest known possible fossil record of aquatic
nematodes consists of tracks dating from the Upper
Precambrian of Australia and Europe (Hantzschel,
1975). Tracks similar to these in the Green River sediments of Lake Uinta (Middle Eocene) (Fig. 2b) were
considered to be made by nematodes (Moussa, 1969).
These more recent tracks occur in the Soldier summit
area near Provo, Utah, and were made during the second lacustrine phase of the Green River sediments.
Lake Uinta was a large freshwater lake that occupied
the Uinta and Piceance Creek Basins in Utah and
Colorado. These tracks could have marked the passage
of fairly large freshwater nematodes moving near the
shore. The early Precambrian tracks could well be
nematodes since it is highly probable that the phylum
dates back to that period when representatives of the
now predominantly aquatic order Araeolaimida
probably existed (Poinar, 1983) and biting midges
262
George O. Poinar, Jr.
(Ceratopogonidae: Diptera) were not present, which
are a group of insects whose legless larvae move in a
fashion similar to nematodes.
The aquatic mermithids represent the earliest and
most frequenty encoutered undisputed nematode fossils.
The earliest of these occurs in 120 – 135 million-year-old
amber from Lebanon (Fig. 12a). It was originally described as occurring in a biting midge (Ceratopogonidae:
Diptera) (Poinar et al., 1994), but according to Art
Borkent (personal communication) the host is actually a
member of the Chironomidae. Since the generic placement of this specimen was based in part on its host family, it is now prudent to transfer the species libani from
Heleidomermis Rubstov into a new genus Cretacimermis
Poinar. Thus, this nematode, which is the oldest definite
fossil nematode, should now be called Cretacimermis
libani Poinar, Acra and Acra (1994). Another aquatic
mermithid fossil species is Heydenius dominicanus from
Dominican amber (Poinar, 1984). Two specimens of this
species were found in close association with an adult
cranefly (Tipulidae: Diptera) and an adult mosquito
(Culicidae: Diptera) (Fig. 12b). Based on the record of
present-day mermithid infections of these two host
groups, and the discovery of a female mosquito with a
mermithid still coiled in its abdomen in Dominican amber, it can be assumed that H. dominicanus was a parasite of the adult mosquito. Such rare fossils show the antiquity of some parasitic associations.
B. Habitats of Freshwater Nematodes
Nematodes constitute an important and significant
portion of the zoobenthic community of freshwater
habitats. This community has been separated by benthic
ecologists into three size categories (Strayer, 1985).
Animals retained on sieves with a mesh ranging from
200 to 2000 m compose the macrofauna, those that
are retained on sieves with a mesh ranging from 40 to
200 m construct the meiofauna, and those that pass
through a 40-m mesh sieve make up the microfauna.
Freshwater nematodes covered in the present chapter
could be considered as belonging to all groups; the large
free-living stages of the Mermithidae would be considered macrofauna, the majority of the adults and juveniles of the other groups would fall under meiofauna,
and the young juveniles and eggs of many forms would
fall into the microfaunal category. Thus, for nematodes
in general, these size categories are little used, although
they are helpful if only certain stages are being studied.
Nematodes constituted 60% of all the benthic
metazoans in Mirror Lake, New Hampshire, and are
probably the most abundant benthic animals in freshwater (Strayer, 1985). In Mirror Lake, nematode abundance showed a distinct depth distribution, with a
strong minimum at 7.5 m. More than 90% of the nematode biomass at 7.5 m was composed of species of
the large predatory nematodes belonging to the genus
Mononchus. Strayer (1985) also reported that 63% of
the nematodes in Mirror Lake lived in the top 2 cm of
sediment, while only 18% penetrated to a depth greater
than 4 cm. Species of the genera Monhystera and Ethmolaimus, which constituted 55% of the nematodes in
Mirror Lake, were found in the benthic zone. Although
the nematodes outranked all other animals in abundance (constituting 59% of all metazoan individuals in
the benthos), they still represented only 1% of the
zoobenthic biomass. Nematodes normally contribute
between 1 and 15% of the zoobenthic biomass in lakes,
and other studies show that they constitute from 40 to
80% of all meiobenthic animals (Holopainen and Paasivirta, 1977; Oden, 1979; Nalepa and Quigley, 1983).
Nematodes that occur in freshwater have traditionally been called free-living nematodes. The term freeliving is an ecological ranking, which means that these
nematodes have no symbiotic association with multicellular plants or animals. Such nematodes obtain
nourishment from bacteria, algae, protozoa, and other
unicellular organisms. Nematodes with this type of nutrition are best referred to as microbotrophic, although
they have been called saprophytic, saprophagous, bacteriophagous, microbivorous, and microphagous. Such
nematodes act as secondary consumers, feeding on bacteria and fungi. They may serve to maintain these microbial populations, although many of the free-living
forms will die if bacterial populations reach high numbers, which is why most of the normal aquatic nematodes are absent from areas high in sewage.
The term free-living, however, is also used to refer
to a particular stage in the life cycle of a nematode —
often a stage in the life cycle of a plant-parasitic or animal-parasitic nematode — when it has either ended or
not yet begun its parasitic relationship. Examples in the
freshwater habitat are the mermithids. The egg, infective second-stage juvenile, postparasitic juvenile, and
adult mermithids are free-living and nonfeeding. Nourishment is obtained from the hemocoel of insects only
during a portion of their juvenile development. Thus,
free-living in the present work refers to those nematodes that have some developmental stages occurring in
freshwater, irrespective of their nutritional requirements or their status at other developmental stages.
The habitats of freshwater nematodes vary as do
the categories of freshwater resources. Two important
requirements for freshwater nematodes to complete
their development are continuation of the water source
and availability of food and oxygen. Many freshwater
nematodes feed on microorganisms that flourish in the
gyttja, an organic deposit composed of excretory
9. Nematoda and Nematomorpha
material from benthic animals. It is most obvious in
lakes and ponds, where it is often combined with decomposing plant debris. This sediment increases in density with increasing depth. Core samples are used for
quantitative measurements of zoobenthos in the gyttja
(Strayer, 1985).
Annual productivity of freshwater nematodes in an
Austrian alpine lake was studied by Bretschko (1973).
In the deeper benthos, below 20 m, there were 2 – 4
annual generations with the total yield of nematode
biomass estimated at 66 kg per year. It is interesting
that the production was higher in shallow water and
during the winter when the lake was covered with ice.
For example, Tobrilus grandipapellatus was collected
at populations of 235,000 individuals / m2 in the winter,
but only 60,000 individuals / m2 in the summer. This
could be related to the availability of oxygen and density of micribial populations.
Nematodes of all environments, including freshwater, are among the most numerous animals feeding on
both primary decomposers such as bacteria and fungi,
as well as primary producers such as algae and higher
plants (Nicholas, 1984). The significance of nematodes
in the general economy of the ecosystem is not known,
and attempts to estimate their contributions to energy
flow have been made only with terrestrial and marine
forms. Such estimates have used the equation:
CPREU
where C is the consumption, P the production, R the
respiration, E the ejecta (feces), and U is excretion.
Although freshwater nematodes have been little
studied, estuarine forms have been examined in detail
(Warwick and Price, 1979). Some 40 species of
TABLE I
263
nematodes occur on a mud flat in the Lynher Estuary in
southern Britain. The population varies from 8 – 9
106 individuals / m2 in winter to nearly 23 106 individuals / m2 in late spring (15% of the macrofauna).
Warwick and Price (1979) calculated a total annual
oxygen (O2) consumption of 28 liter O2 /m2 per year,
which is equivalent to some 11 g of metabolized carbon (C). The annual production was calculated at
about 6.6 g C / m2 per year. Previous calculations with
the soil nematode Caenorhabditis briggsae showed that
the conversion of bacteria into nematode tissue varied
from 13 to 20% (Nicholas, 1984).
The question of nematodes living under anoxic
conditions has been addressed by several workers. In
deep lakes, the hypolimnion may become anaerobic for
certain periods, resulting in a reduction of nematode
fauna, but not eradication. In Lake Tiberias, in Israel,
Por and Moary (1968) found large numbers of Eudorylaimus andrassyi at a depth of 43 m during the winter oxygen-free period.
In the Neusiedlersee in Austria, Tobrilus gracilus
occurs in muddy anaerobic zones beyond the limits of
emergent vegetation (Schiemer, 1978). In his study of
the nematodes in Mirror Lake, Strayer (1985) found
most species in the littoral zone, but species of Ethmolaimus, Monhystera, and unidentified Tylenchidae were
highly abundant in the anaerobic sediments at a depth
of 10.5 m.
Extreme habitats for freshwater nematodes include high-temperature hot springs. Table I lists some
freshwater nematodes collected at unusually high
temperatures. The listing of 61.3°C for Aphelenchoides sp. is not only a record for nematodes, but for
all metazoan life forms!
Survival of Freshwater Nematodes in Hot Water Springs at High Temperatures
Nematode species
Temperature (0C)
Location
Reference
Aphelenchoides sp.
Aphelenchoides sp.
Aphelenchus sp.
Doryalimus atratus Linstow
Dorylaimus atratus Linstow
[D. thermae Cobb]
Dorylaimus atratus Linstow
[D. thermae Cobb]
Euchromadora striata (Eberth)
Monhystera ocellata Bütschli
Monhystera gerlachii Meyl
Plectus sp.
Rhabdolaimus brachyuris Meyl
Theristus pertenuis Bresslau and
Schuurmans Stekhoven
Tylocephalus sp.
61.3
35.0
57.6
47.0
40.0
New Zealand
New Zealand
Chile
Italy
Wyoming, USA
Rahm (1937)
Winterbourn and Brown (1967)
Rahm (1937)
Issel (1906)
Hoeppli (1926)
53.0
Wyoming, USA
52.0
52.0
52.0
57.6
52.0
52.0
Italy
Italy
Italy
Chile
Italy
Italy
Cobb, in Hoeppli
(1926)
Meyl (1954)
Meyl (1954)
Meyl (1954)
Rahm (1937)
Meyl (1954)
Meyl (1954)
45.3
New Zealand
Winterbourn and Brown (1967)
264
George O. Poinar, Jr.
Other specialized habitats encompass brackish and
estuarine waters, inland saline lakes, cave streams, and
associations with specific aquatic plants. “Artificial water sources” include canals, waste water, tap water,
wells, filter beds of sewage treatment plants, and temporary water sources. Nematodes found in these habitats often do not have any relation to true freshwater
forms or a freshwater habitat, and the majority are not
treated in this work.
A unique cave habitat has resulted in what appears
to be the first aquatic nematode specialized for survival
in hydrogen sulfide-rich thermo-mineral waters. These
conditions are found in the Movile cave, located in a
limestone plateau in Southern Dobrogea, Romania
(Poinar and Sarbu, 1994). The nematode, Chronogaster troglodytes, lives in floating microbial mats comprised primarily of mycelia of fungi belonging to the
class Oomycetes and various bacterial populations. Selection for this habitat could have undergone millions
of years since a Miocene age for the origin of Movile
cave has been suggested. Also, while members of the
genus Chronogaster are mostly aquatic, they, like many
freshwater nematodes, are quite broad in their selection
of habitat. Other species of Chronogaster have been
FIGURE 3
collected from sandy or sandy loam soils, clay soils,
pasture soils and even forest soils. In fact one species,
C. africana is mentioned as occurring in multiple
aquatic and terrestrial habitats (Heynes and Coomans,
1980). Such plasticity was probably an important feature in the specialization of C. trogoldytes in its unique
underground cave habitat. A list of nematodes recovered from caves and subterranean waters is presented
by Poinar and Sarbu (1994).
Previous workers have attempted to separate the
freshwater-dependent nematodes from the facultative
forms. Micoletzky (1922) distinguished between completely aquatic, principally aquatic, amphibious, and
principally terrestrial species; while more recently, Jacobs (1984) presented an ecological classification with
stenohygrophilic nematodes representing the strictly
aquatic forms and enhygrophilic nematodes representing the semi-aquatic forms. The strictly aquatic or
stenohygrophilic taxa covered here occur in many microhabitats (Fig. 3). As examples, periphytic forms live
among plant roots, bryophilic taxa associate with moss
and liverworts, endobenthic species thrive within the
sediment and bottom debris, epibenthic groups live on
the surface of the bottom, haptobenthic forms exist on
Scene of the bottom of a typical fast-flowing stream showing several nematode microhabitats. (A)
Stream bed containing endobenthic forms; (B) bed surface containing epibenthic groups; (C) rock surface containing haptobenthic groups; (D) Nostoc algal pads containing mermithids that parasitize Cricotopus midges
(Chironomidae) living inside the algae.
9. Nematoda and Nematomorpha
the surface of submerged rocks, debris, and aquatic
plants, and planktonic taxa live continuously in the
water column. The latter group is poorly known and
mainly found in turbid waters where their occurrence
may represent more of a dispersal than a habitat mode.
Many genera of freshwater stenobygrophilic nematodes are widely dispersed, occurring on more than one
continent. Some species are eurytopic, occurring in diverse habitats, while others are stenotopic and restricted to a few specialized habitats.
Freshwater nematodes have been considered as indicators of water pollution (Zullini, 1976). Heavy metals
and other pollutants settle and are taken up by organic
matter in the sediments. Nematodes ingest this material
and those that are sensitive to the pollutants may die.
Representatives of the Chromodorida are apparently
more sensitive to pollution that those of the Rhabditida
(Zullini, 1976). In extremely polluted waters, only the
Rhabditoidea may be quite abundant. A two-year study
sampling nematodes along two streams in Indiana indicated that an analysis of the benthic nematode community structure could be useful in evaluating disturbance
to aquatic habitats (Ferris and Ferris, 1972).
C. Enemies
Perhaps the greatest enemies of freshwater nematodes are other predaceous nematodes. Representatives
of the Mononchida, Dorylaimida, and Enoplida are
known to attack and devour a range of small invertebrates in their surroundings. The crayfish Pacifastacus
leniusculus was observed to feed occasionally on nematodes (Flint, 1976). The freshwater, rhabdocoel turbellarian, Microstomum feeds on nematodes (Stirewalt,
1937), as will the freshwater nemertean worm, Prostoma (Coe, 1937). The role of disease organisms in
regulating populations of freshwater nematodes is not
known, but the author has frequently collected specimens of Tobrilus infested with what appeared to be
microsporidian spores (Fig. 9c). Other records of probable protozoan diseases in freshwater nematodes have
been reported in Chromadora, Dorylaimus, Ironus,
Monhystera, Plectus, Theristus, Trilobus, Tripyla,
Prodesmodora, Achromadora, Paraphanolaimus, and
Desmolaimus (Poinar and Hess, 1988).
V. COLLECTING AND REARING TECHNIQUES
265
are present in the ecosystem. The latter method is used
to evaluate the components of a population, the proportion of each in the environment, and the dynamics
of the population over time.
Qualitative methods for sampling freshwater nematodes depend on the water flow and the zone to be sampled. In sampling nematodes from beds of fast-flowing
streams, a net, an instrument to turn over rocks and debris on the stream bed, and a set of sieves, are necessary
(Fig. 4a). The net is first anchored with the opening facing upstream. A pick or geologist’s hammer is used to
disturb the stream bed slightly upstream from the net
(Fig. 4b). The stream washes nematodes and other debris from the disturbed area into the net. The contents
of the net are then placed in enamel collecting pans with
water and set aside for extraction. Nets can be pulled
over aquatic plants to obtain nematodes living on their
surfaces. The type of nematode being collected will determine the mesh size of the net selected. It is recommended that, in order to obtain a fair sample of all nematodes, a mesh size no larger than 40 m be used.
A number of devices have been devised to collect
freshwater nematodes (Filipjev and Schuurmans
Stekhoven, 1959). These include dredges, wormnets,
bottom catchers, mudsuckers, swab or sledge trawls,
and plankton nets. Their use depends on the type of material desired. For sluggish or stationary water sources,
samples of the bottom (mud, sand) can be placed in a
water-filled container (Fig. 4c) which is then shaken, the
suspension allowed to settle for 3 – 4 sec, and the supernatant then poured into a second container for extraction. Rocks and other submerged material can be placed
in buckets and the nematodes washed off with a water
spray or by vigorous shaking or brushing.
Quantitative sampling methods involve the use of
augers or probes to remove core samples of measurable
sizes from the beds (Strayer, 1985). Other quantitative
methods can be devised on the basis of need and the
desired biotype to be investigated. Techniques applied
to marine nematodes may also be suitable for freshwater forms. For further information, see Hulings and
Gray (1971), Holme and McIntyre (1971), and Downing and Rigler (1984). It should be remembered that
any estimate of the nematode population depends on
the type of sample taken. Thus, a biased sample will
produce a biased estimate of the nematode population.
Also, it should be noted that nematodes are sensitive
animals and all sampling and extraction techniques
should be as gentle as possible.
A. Sampling Methods
There are two basic types of sampling: qualitative
and quantitative. The former method is used in preliminary studies to determine which kinds of nematodes
B. Extraction
The process of extraction involves removing nematodes from samples collected in various freshwater
266
George O. Poinar, Jr.
FIGURE 4
(a) Equipment needed to sample freshwater nematodes. A, pail; B, set of gradated sieves; C, net;
D, pick; E, squeeze bottle; F, shovel, G, geologist’s hammer. (Hip boots are not shown.) (b) With the geologist’s
hammer, the stream bed is disturbed upstream from the positioned net. (c) With a shovel, samples of the
stream bed are removed together with roots of aquatic plants. (d) Large nematodes (mermithids and dorylaimids) and debris are transferred from the net into an enamel examination pan.
habitats (Figs. 4d, 5a–d). The basic principles of nematode extraction are based on the fact that most nematodes have a specific gravity between 1.10 and 1.14,
thus they will slowly sink in freshwater. They are denser
than fine clay particles, but lighter than sand. The extraction methods used for soil and plant-parasitic nematodes are wholly adequate for freshwater forms.
Although the majority of freshwater nematodes are
too small to view with the naked eye, the free-living
stages of many aquatic mermithids are large enough to
be handpicked from the samples (Fig. 5c). They can be
carefully lifted with a fine needle containing an Lshaped bend at the tip or with a pair of fine forceps
(Fig. 5d). Other mermithids that are small enough to
9. Nematoda and Nematomorpha
FIGURE 5
(a) Nematodes and debris collected from the stream bed are passed through a series of three sieves
with openings ranging from 0.4 – 0.04 mm across. (b) Nematodes are washed off the surface of the final screen
with a spray of water from the plastic squeeze bottle. (c) Free-living stages of mermithids are large enough to
be seen with the naked eye and can be picked up with forceps. (d) Larger nematodes can be transferred directly
to a small vial for later killing and fixing.
267
268
George O. Poinar, Jr.
escape visual spotting can be collected with the extraction processes.
One of the oldest and simplest methods of extracting nematodes from a wide range of samples is the
Cobb sieving and gravity method, or a modification of
the latter. This consists of making a water suspension
of the nematodes and pouring it through a series of
screens of different mesh sizes to collect the nematodes.
The suspension is made by placing the samples in a pail
or other suitably large container (Fig. 4c), adding
2 – 3 L of water (preferably from the original source),
allowing the mixture to set for 10 – 20 sec to allow the
heavier particles to sink, and then pouring the suspension containing the nematodes through a series of
screens (Fig. 5a). If extraneous large debris (wood particles, rocks) is present, the samples should first be
poured through a 20 – 40 mesh sieve (openings are
0.840 – 0.350 mm). The sizes of sieves selected will, of
course, influence the size range of nematodes extracted.
If a sample of all available nematodes is desired, then a
final fine screen of 400 – 500 mesh (openings are
0.035 – 0.026 mm) can be used. The problem with the
fine screens is that they become easily clogged with debris. Such screens should be tapped rapidly and gently
on the under surface with the fingers to assist passage
of the water. The nematodes, together with debris, will
be trapped on the surface of the 400- or 500-mesh
sieve. Both can be removed by directing a small jet of
water on the back of the sieve and washing the contents into a separate container (Fig. 5b). Certain biases
are inherent in sieving extraction methods (Hummon,
1981). Since they may be unsuitable for quantitative
studies (Viglierchio and Schmidt, 1983), the more
tedious method of sorting through unsieved samples
under a dissecting microscope may be desirable
(Strayer, 1985).
The final step consists of separating the nematodes
from the fine debris. For this, the Baermann funnel
method, or a modification thereof, can be used. All that
is needed is a funnel with a piece of rubber tubing attached to the stem. The tube should be closed with a
screw clamp. The nematode – debris mixture is placed
on a fine cloth or facial tissue, which is supported by a
piece of screen held in the top, wide portion of the funnel. Water is slowly added to the funnel until the nematode – debris mixture is just covered. The clamp should
be opened to eliminate the air trapped in the funnel
stem and to allow a continuous column of water to
flow into the funnel. The nematodes crawl through the
debris and facial tissue, drop through the screen, and
settle at the base of the funnel stem, where they can be
drawn off by releasing the clamp. The nematodes
should be drawn off every 6 h; the apparatus can be
operated in this fashion for several days. Then the
nematodes can be counted or handpicked under a dissecting microscope.
A more recently devised method of isolating nematodes from samples involves the principle of increasing
the specific gravity of the solution to make the nematodes float to the surface. Substances such as sugar,
salt, or Ludox can be added to the nematode – water
mixture in the centrifugal-flotation and sugar-flotation
techniques (Ayoub, 1977). If sugar is used, then 673 g
added to 1 L of water will result in a specific gravity of
1.18, which is greater than that of nematodes (causing
them to float on the surface). Detailed instructions for
these and other methods of extracting nematodes from
soil and plant samples are presented by Ayoub (1977)
and Nichols (1979).
Quantitative methods of extraction for monitoring
nematode populations over a period of time often involve the use of elutriators, which use an upcurrent of
water to separate nematodes from the medium. Such
methods are rather elaborate and are discussed by
Southey (1978).
C. Fixing and Mounting
After the nematodes have been extracted, they
should be held in water (preferably water from their
original collecting source) prior to fixation. The nematodes should be killed before being placed in fixative;
otherwise they become distorted and difficult to examine. They are most easily killed with heat, which also
tends to relax and extend them. Too much heat will
destroy the internal tissues, but good results can be
obtained by pouring hot water (60 – 70°C) over the
nematodes. They should then be transferred to the
fixative as soon as possible. The best fixative for
nematodes is TAF (7 mL 40% formalin, 2 mL triethanolamine, and 91 mL distilled water). However,
3 – 5% formalin or 70% ethanol can also be used if
TAF is not available. Some distortion may occur with
ethanol, however.
The nematodes should be fixed for at least 2 – 3
days before transfer to glycerin for mounting on microscope slides. Nematodes are most easily conveyed to
glycerin by the evaporation method, which requires little handling. Fixed specimens are transferred to a dish
containing a solution of 70 mL ethanol (95%), 5 mL
glycerol, and 25 mL water. The dish is partly covered
for the first 3 days to allow the alcohol and water to
evaporate at a slow rate; then the cover is removed for
the next 14 days. Finally, the containers (now with
mostly glycerin) are placed in a desiccator or an oven
(35°C) for another 2 weeks to drive out the remaining
water. The nematodes will then be in a relatively pure
solution of glycerin and can be mounted directly on
9. Nematoda and Nematomorpha
microscope slides. Such slides are termed permanent
since they can be kept for years for continuous study.
Eventually, the nematode tissues tend to become transparent. Temporary slides are made by mounting the nematodes directly after fixation in the fixing solution
(formalin or alcohol). They may last several months if
the ringing seal is tight.
The following steps can be taken to make permanent slides with specimens transferred to glycerin
(modified from Poinar, 1983):
1. With a small pointed instrument (dental pulp
canal file, small insect pin mounted on a
wooden splint, needle), transfer the nematode(s)
to a small drop of glycerin placed in the center
of a microscope slide.
2. Push the nematode to the bottom of the drop.
Then add at three equidistant points around the
nematode, small supports (coverslip pieces,
wire) with a width equal to or slightly wider
than that of the nematode. Push these also to
the bottom of the drop.
3. Place a cover slip over the drop and lower it
slowly at a 30° angle so that air bubbles will
not become entrapped in the glycerin as the
slide touches the mounting medium.
4. Add more glycerin, if needed, by placing a small
drop at the edge of the coverslip and allowing it
to move under the glass and spread throughout.
If the coverslip is floating on glycerin, then remove the excess by blotting it with a moistened
(water) piece of tissue.
5. Carefully seal the edges of the coverslip with a
ringing compound, such as nail polish or Turtox
slide ringing cement (General Biological Supply
House, Chicago, II).
6. Label the slide with information regarding the
date, locality, and collector.
D. Culturing
Some free-living, microbotrophic freshwater nematodes can be cultured in the laboratory if they are presented with a growing colony of their preferred food
(i.e., bacterial or algal species) and maintained under
environmental conditions (temperature, oxygen) comparable to those present at the collecting site. Nuttycombe (1937) successfully cultured several species of
freshwater nematodes using a wheat grain infusion
method. Between 200 and 300 grains of wheat seeds
were added to 250 mL of spring water in a flask. This
mixture and a separate flask of pure spring water were
heated to a boil and allowed to cool. The spring water
was poured into 200 mL Petri dishes with 3 – 4 grains
269
of the boiled wheat. The dishes with the wheat seeds
were allowed to stand for 2 – 3 days before the nematodes were added. Microorganisms brought in with the
nematodes grew on the wheat seeds, and the microbotrophic nematodes fed on the microbes.
Postparasitic juveniles of freshwater mermithids
can be held in the laboratory until they mature for
identification. Care should be taken so that both the
temperature and the amount of dissolved oxygen parallel those of the collection site. Periodic transfers to new
containers with freshwater may be necessary to prevent
the buildup of fungi which will destroy the nematodes.
Petersen (1972) devised a method of culturing a mosquito mermithid parasite (Romanomermis culicivorax)
through its entire life cycle. The mosquito chosen as a
laboratory host was Culex pipiens quinquefasciatus
(Say), because it could be reared continuously in
crowded conditions, was easily maintained in colony,
and was highly susceptible to the nematode. Postparasitic juvenile mermithids were placed in wet sand
within plastic trays (36 25 100 cm). The postparasites molted to the adult stage, mated, and oviposited
in the sand. The eggs, which embryonated in the moist
sand, could be maintained in their resting state for several months. When the trays were flooded with water,
the infective stages emerged from the sand and were
capable of infecting newly hatched mosquito larvae.
The infected insect larvae were kept in a separate container and fed. By the time the mosquitoes were ready
to pupate, the nematodes had completed their development and started to emerge. The emerging postparasitic
juveniles were then transferred to wet sand to continue
the cycle. Adequate numbers of mermithids were available for both scientific investigations and biologic
control studies with this rearing method.
VI. IDENTIFICATION
Once the specimens are mounted, they can be examined and identified with a compound microscope.
Because the identification of nematodes involves the
use of many fine details, a microscope equipped with
oil immersion should be employed (1000 ). The use of
dioscopic illumination after Nomarski or differential
interference contrast better reveals minute features of
the cuticle (such as the amphid structure). If possible,
living or just-killed nematodes should also be examined
under the microscope. Their movements can be reduced by removing water from under the coverslip and
compressing the nematode between the microscope
slide and coverslip. Certain characters are much clearer
on living specimens and there is a certain thrill
in watching these creatures move under a microscope
270
George O. Poinar, Jr.
that is absent with preserved material. However,
measurements can best be made on fixed nematodes.
In preparing the following taxonomic key, the following works dealing with freshwater nematodes were
consulted: Cobb (1914); Chitwood and Allen (1959);
Ferris et al. (1973); Tarjan et al. (1977); Pennak (1978,
1989); Esser et al. (1985); Esser and Buckingham
(1987); Jacobs (1984); Tsalolikhin (1983); Gerber and
Smart (1987). The first five references also have keys
which can be consulted by the interested reader for
cross-reference. For descriptions of various genera and
a listing of species, Goodey (1963) is still very helpful,
although now out of date.
Some 66 genera of freshwater nematodes are included in the present key. Between 300 and 500 species
of nematodes are probably included within these 66
genera. Only those genera that contain species that are
obligately aquatic (live nowhere else) and have been reported from North America are included. Over 95% of
the aquatic nematodes encountered should be included
in the following key.
A. Taxonomic Key to Families and Selected Genera
of Freshwater Nematoda
The following key pertains to the families and selected genera of freshwater nematodes and freeliving
stages of parasitic nematodes found in freshwater habitats of North America. Following this is a separate key
to the genera of aquatic Mermithidae.
1a.
Elongated and threadlike, generally over 1 cm in length and usually observable with the naked eye; found on the bottom surface or
several centimeters within the bed (Figs. 5c and 10a, b) ....................................................................................................................2
lb.
Not elongate, generally under I cm in length and observable only with a lens (Figs. 2c and 11c); found on all exposed surfaces
(rocks, on or in plants) as well as on the bottom surface and within the bed ......................................................................................3
2a (1a).
Forms usually long (6 cm or more in length), leathery body wall, range in color from light brown to black (Fig. 13a) ........................
Hairworms (phylum Nematomorpha)
2b.
Forms usually smaller (under 6 cm in length), body wall fragile, range in color from white to rose, green and yellow (Fig. 5c) ...........
Mermithidae (see Section VI.B.)
3a (1b).
Head of nematode bearing a stylet (Figs. 8b and 9b) ..........................................................................................................................4
3b.
Teeth and other armature may be present, but a stylet is absent (Figs. 7a, b and 8a) ........................................................................16
4a (3a).
Stylet knobs usually absent (Fig. 9b); if present, then pharynx lacking a valvated metacorpus; pharynx narrow anterior and thickened posterior at junction with intestine (Fig. 2a); valvated metacorpus absent ...........................................................Dorylaimida 11
4b.
Stylet knobs usually present (Fig. 8b); pharynx composed of a valvated metacorpus followed by a slender isthmus and basal glandular bulb leading into the intestine (Figs. 8b, c) ....................................................................................................................................5
5a (4b).
Dorsal gland outlet in precorpus; metacorpus moderate in size (Fig. 8b) (less than three-fourths of body width ..............Tylenchida 8
5b.
Dorsal gland outlet in metacorpus; metacorpus large (three-fourths of body width or more) (Fig. 8c) .................................................
Aphelenchida .....................................................................................................................................................................................6
6a(5b).
Stylet usually with faint or inconspicuous knobs (Fig. 8c); tail tip usually pointed; males lacking bursa and gubernaculum.................
Aphelenchoididae ...............................................................................................................................................................................7
6b.
Stylet without knobs; tail tip usually rounded; males with a bursa (Fig. 6c) and gubernaculum ...........................................................
Aphelenchidae Aphelenchus Bastian, 1865
7a (6a).
Tail shape elongate, filiform .................................................................................................................................Seinura Fuchs, 1931
7b
Tail shape short, conical ........................................................................................................................Aphelenchoides Fischer, 1894
8a(5b).
Head bearing distinct setae (Fig. 7a) .......................................................................................Atylenchidae Atylenchus Cobb, 1913
8b
Head without setae ...........................................................................................................................................................................9
9a (8b).
Procorpus fused with large, oval metacorpus; cuticle strongly annulated; adult female with cuticular sheath and well-developed
stylet (Fig. 8b) ........................................................................................................Criconematidae Hemicycliophora de Man, 1921
9b.
Procorpus distinct and narrow before reaching the expanded metacorpus, cuticle not strongly annulated; adult female without cuticular sheath; styles may or may not be well developed.............................................................................................Tylenchidae 10
10a (9b).
Large distinct stylet, ovaries paired, amphidelphic (Fig. 6a); tail tapering but not filiform..........................................Hirschmanniella
Luc and Goodey, 1963
10b.
Stylet small and inconspicuous, ovary single, prodelphic; tail filiform ............................................................Tylenchus Bastian, 1865
11a (4a).
Pharynx gradually widens toward basal bulb (Fig. 2a) .....................................................................................................................12
FIGURE 6
(a) Midbody region of a female Pellioditis sp. (Rhabditidae) showing the vagina (arrow) and
paired opposite uteri filled with developing eggs. Note copulation deposit surrounding the vulva opening. (b)
Longitudinal striations lining the cuticle of Dorylaimus sp. (Dorylaimidae). (c) Tail of male Pellioditis (Rhabditidae) showing spicule (S), gubernaculum (G), bursa (B), and bursal papillae or rays (arrows). (d) Tail of an
aquatic nematode showing caudal gland mucous being emitted from the terminus (arrow).
271
FIGURE 7
(a) Anterior end of an aquatic nematode showing a narrow tube-shaped stoma (arrow) and
cuticular bristles or setae. (b) Large oval, cup-shaped stoma of a predatory aquatic nematode. Note dorsal
tooth (arrow) on stomal wall. (c) Nematode bearing a medium-sized, circular amphid. (d) Nematode bearing a
large, spiral-shaped amphid.
272
FIGURE 8 (a) Pharyngeal region of Pellioditis (Rhabditidae) showing tubular stoma (S), corpus (C), metacorpus (M), isthmus (I), and basal bulb (B). Arrow shows valve in basal bulb. (b) Outer ensheathing cuticle of
Hemicycliophora (Hemicycliophoridae). Note long stylet (S) with prominent basal knobs (arrow) and valvated
metacorpus (M). (c) Anterior of Aphelenchoides sp. (Aphelenchoididae) showing small stylet lacking knobs
(arrow) and large metacorpus (M) with valve. (d) Outstretched ovary (arrow shows tip). (e) Reflexed ovary
(arrow shows point of reflexion).
273
274
George O. Poinar, Jr.
11b.
Pharynx abruptly widens to form the basal bulb which contains a valvular chamber (Fig. 8a) ......................................Campydoridae
Aulolaimoides Micoletzky, 1915
12a (11a). Head bearing a narrow protrusible mural spear that is symmetrically pointed at tip ..............Nygolaimidae Nygolaimus Cobb, 1913
12b.
Head bearing a thick protrusible axial spear that is asymmetrically pointed at tip (sloped on one side) (Fig. 9b) .................................
Dorylaimidae ...................................................................................................................................................................................13
13a (12b). Stomal area bearing distinct teeth or denticles around the tip of the stylet .......................................................................................14
13b.
Stomal area lacking distinct teeth or denticles in the stylet tip region ...............................................................................................15
14a (13a). Four large teeth together with mural denticles present ............................................................................Paractinolaimus Meyl, 1957.
14b.
Four large teeth only present ........................................................................................................................Actinolaimus Cobb, 1913
15a (13b). Cuticle thick and longitudinally ridged (Fig. 9b) .......................................................................................Dorylaimus Dujardin, 1845
15b.
Cuticle smooth ..................................................................................................................................Mesodorylaimus Andrassy, 1959
16a (3b).
Amphids minute and borne on the lateral lips, excretory duct cuticularized (canal lined with a fine layer of cuticle); caudal glands
absent, conspicuous cephalic setae absent, bursa present or absent ................................................................................................ 17
16b.
Amphids usually enlarged and located on the neck region, excretory duct rarely cuticularized (except in Plectidae); head often with
conspicuous setae; bursa usually absent. ..........................................................................................................................................21
17a (16a). Pharynx with an expanded median bulb (metacorpus) containing a longitudinal valve and a glandular nonvalvated basal bulb
Diplogasteridae ...................................................................................................................................... Rhabditolaimus Fuchs, 1915
17b.
Pharynx may or may not contain a metacorpus, but if it does, it does not contain a valve; basal pharyngeal bulb with or without a
basal valve........................................................................................................................................................................................18
18a(17b).
Pharynx elongate and lacking a valve in the basal bulb or bulb area, slender; slow moving forms (Fig. 9d) ....................Daubayliidae
Daubaylia Chitwood and Chitwood, 1934
18b.
Pharynx normal in length, with a basal bulb containing a valve (Fig. 8a); stouter, quick-moving forms ...........................................19
19a(18b).
Stoma with rhabdions separate, female with a single ovary, bursa absent ..................................................................Cephalobidae 20
19b.
Stoma with rhabdions fused, cylindrical (Fig. 8a), female with a single or paired ovaries, bursa usually present (Fig. 6c) ....................
Rhabditidae [Most of the genera in this large family are soil forms, some of which appear from time to time in aquatic habitats.
Mesorhabditis (Osche, 1952), which has a single ovary and a bursa, occurs ;n sewage bed and run-off water. Pellioditis (Doughtery,
1953), which has paired ovaries also occurs in aquatic habitats.]
20a (19a). Lateral fields reach to tail tip; female tail rounded .......................................................................................Cephalobus Bastian, 1865
20b.
Lateral fields end at the phasmids; female tail usually pointed...................................................................Eucephalobus Steiner, 1936
21a (16b). Amphids spiral (Fig. 7d), loop-like, stirrup-shaped or circular (Fig. 7c); pharynx usually with a basal bulb (Fig. 8a) .......................22
21b.
Amphids pore-like to pocket-like, pharynx usually without a distinct basal bulb .............................................................................41
22a (21a). Ovary usually outstretched (reflexed in Prismatolaimus) (Fig. 8d), amphids usually circular (Fig. 7c) but may be slitlike ....................
Monhysterida...................................................................................................................................................................................23
22b.
Ovaries reflexed (Fig. 8e) (except in Odontolaimus which also has circular amphids), amphids variable (including circular)...........25
23a (22a). Amphids slit-like, faint; stoma with denticulated cushions (padded areas covered with minute projections).........................................
Prismatolaimus de Man, 1880
23b.
Amphids circular, distinct, stoma lacking denticulated cushions .......................................................................................................24
24a (23b). Stoma shallow; pharynx without distinct basal bulb (as in Fig. 2a) .............................................................Monhystera Bastian, 1865
24b.
Stoma elongate; pharynx with distinct basal bulb (as in Fig. 8) ....................................................................Monhystrella Cobb, 1918
25a (22b). Caudal glands and spinneret present (Fig. 2d); stoma usually armed with teeth (Fig. 7b); knobs, bristles (Fig. 7a), or punctations
usually present on cuticle .........................................................................................................................................Chromadorida 26
25b.
Caudal glands and spinneret present or absent; stoma may or may not be armed with teeth; bristles sometimes present on cuticle,especially around head ..................................................................................................................................................Araeolaimida 33
26a (25a). Amphids circular (Fig. 7c); cuticular punctations minute ..................................................Microlaimidae Microlaimus de Man, 1880
26b.
Amphids spiral (Fig. 7d), kidney shaped or circular; cuticular punctations coarse ............................................................................27
27a (26b). Amphids circular (as in Fig. 7c) or spiral (as in Fig. 7d), pharyngeal – intestinal junction large, tri-radiate..............Cyatholaimidae 29
FIGURE 9 (a) Posterior portion of a postparasitic juvenile mermithid (Mermithidae) in the process of
molting. Note shedding of third- (3) and fourth-stage (4) cuticles. A, adult. (b) Head of Dorylaimus sp. (Dorylaimidae) showing base of lip region (arrows) and stylet lacking basal knobs. (c) Tail region of a Tobrilus sp.
(Tobrilidae) showing spores of a microsporidian parasite filling the hypodermal tissue (arrows). (d) Adults of
Daubaylia (Daubayliidae) inside an aquatic snail.
275
276
George O. Poinar, Jr.
27b.
Amphids spiral or kidney shaped, pharyngeal – intestinal junction small, not tri-radiate .........................................Chromadoridae 28
28a (27b).
Amphids represented as a broad transverse slit; five or more preanal genital papillae in male.................Chromadorita Filipjev, 1922
28b.
Amphids represented as a flattened, broken ring; no more than three preanal genital papillae in male .................................................
Punctodora Filipjev, 1930
29a (27a). Amphids circular (Fig. 7c) ................................................................................................................................................................30
29b.
Amphids spiral (Fig. 7d) ...................................................................................................................................................................31
30a (29a). Stoma cuplike; length shorter than neck width ..................................................................................Prodesmodora Micoletzky, 1923
30b.
Stoma tubular (Fig. 7a); length two or more times neck width ...............................................................Odontolaimus de Man, 1880
31a (29b). Stoma tubular (Fig. 7a), there may be three anterior equal teeth in the stoma, but single large dorsal tooth is lacking .........................
Ethmolaimus de Man, 1880
31b.
Stoma cup- or funnel-shaped, armed with a prominent dorsal tooth (as in Fig. 7b) much larger than any other teeth......................32
32a (31b). Amphids located at level of stoma, small subventral teeth absent .................................................Paracyatholaimus Micoletzky, 1924
32b.
Amphids located posterior to stoma, one or two small subventral teeth present .........................................Achromadora Cobb, 1913
33a (25b). Caudal glands absent; stoma funnel-shaped with separate rhabdions ..................................................................Teratocephalidae 34
33b.
Caudal glands present (Fig. 2a) or absent; stoma tubular with rhabdions fused ...............................................................................35
34a (33a). Amphids porelike; cuticular annulation strong (Fig. 8b) ........................................................................Teratocephalus de Man, 1876
34b.
Amphids spiral (Fig. 7d); cuticular annulation weak ........................................................................Euteratocephalus Andrassy, 1958
35a (33b). Ovary outstretched (Fig. 8d); amphids circular.............................................................................................................Axonolaimidae
Cylindrolaimus de Man, 1880
35b.
Ovary reflexed (Fig. 8e) ....................................................................................................................................................................36
36a (35b). Pharynx usually with a basal valvated bulb (Figs. 1 and 8a).............................................................................................................37
36b.
Pharynx with or without a basal bulb, if bulb present, then valve absent .........................................................................................39
37a (36a). Basal bulb of pharynx lacking valves ................................................................................................................Anonchus Cobb, 1913
37b.
Basal bulb of pharynx with valves (Fig. 8a) ......................................................................................................................................38
38a (37b). Pharyngeal basal bulb with postbulbar extension (Fig. 10c) connecting it with the intestine; caudal glands absent ..............................
Chronogaster Cobb, 1913
38b.
Phayrngeal basal bulb without postbulbar extension, abutting the intestine; caudal galnds present (Fig. 1) ..............................Plectus
Bastian, 1865
39a (36b). Basal bulb lacking .......................................................................................................................Bastianiidae Bastiania de Man, 1876
39b.
Basal bulb present ................................................................................................................................................Camacolaimidae 40
40a (39b). Amphids circular (Fig. 7c), basal pharyngeal bulb absent .......................................................................Aphanolaimus de Man, 1880
40b.
Amphids spiral (Fig. 7d), a slight basal pharyngeal bulb present ...................................................Paraphanolaimus Micoletzky, 1923
41a (21b). Stoma usually oval in outline (Fig. 7b), heavily sclerotized and armed with one or more teeth, setae and bristles absent .....................
Mononchidae
41b.
Stoma not in the form of a heavily sclerotized oval cavity, teeth or denticles may be present, setae and bristles may be present
(Fig. 7a)............................................................................................................................................................................................42
42a (41b). Cuticle of head double.............................................................................................................................................Oncholaimidae 43
42b.
Cuticle of head normal .....................................................................................................................................................................44
43a (42a). Setae or bristles present on tail ...............................................................................................................Oncholaimus Dujardin, 1845
43b.
Setae and bristles usually lacking on tail......................................................................................................Mononchulus Cobb, 1918
44a (42b). Stoma heavily sclerotized, cylindrical .................................................................................................................................Ironidae 49
44b.
Stoma not heavily sclerotized, funnel-shaped or tubular...................................................................................................................45
45a (44b). Stoma vestigial and unarmed; pharynx base expanded and set off to form a slight, elongate bulb...................................Alaimidae 46
FIGURE 10
Chronogaster troglodytes Poinar and Sarbu, a specialized aquatic cave nematode from Movile
Cave in Romania. (a) Anterior end with funnel-shaped mouth cavity. (b) Head showing stirrup-shaped amphid
with a circular opening. (c) Basal pharyngeal region showing valve (fleche) in basal bulb and postbulbar extension (arrow) between the basal bulb and intestine. (d) Spermlike bodies (arrow) which occur near the proximal
portion of the female gonad, and the absence of males, suggests that C. troglodytes is hermaphroditic.
277
278
George O. Poinar, Jr.
45b.
Stoma distinct, or if vestigial then armed with an inconspicuous median tooth; pharynx may be expanded at base, but rarely set off
from the remainder in the form of a bulb .........................................................................................................................................47
46a(45a).
Amphids porelike, minute ................................................................................................................................Alaimus de Man, 1880
46b.
Amphids cup-shaped, distinct......................................................................................................................Amphidelus Thorne, 1939
47a(45b).
Base of pharynx expanded to form a valvated bulb; three rodlike thickenings compose posterior part of stoma ..................................
Leptolaimidae .........................................................................................................................................Rhabdolaimus de Man, 1880
47b.
Pharynx cylindrical, not with basal valvated bulb; stoma a simple tube without rodlike thickenings ..............................Tripylidae 48
48a(47b).
Three lips; amphids porelike, minute; stoma cylindrical .....................................................................................Tripyla Bastian, 1865
48b.
Six lips; amphids cup-shaped, distinct; stoma funnel-shaped ..........................................................................Tobrilus Andrassy, 1959
49a(44a).
Stoma tubular, with two minute teeth at the base; excretory pore opening posterior to head .......................Cryptonchus Cobb, 1913
49b.
Stoma tubular, with three anterior hooklike teeth; excretory pore opening in head area ......................................Ironus Bastian, 1865
50a(41a).
Pharyngeal-intestinal junction tuberculate; dorsal tooth on stoma pointing posteriorally...............................Anatonchus Cobb, 1916
50b.
Pharyngeal-intestinal junction not tuberculate; dorsal tooth on stoma pointing anteriorally ............................................................51
51a (50b). Ventral stomatal ridge with longitudinal row of denticles ..............................................................................Prionchulus Cobb, 1916
51b.
Ventral stomatal ridge absent, or if present then unarmed ...........................................................................Mononchus Bastian, 1865
B. Taxonomic Key to Extant Genera
of Freshwater Mermithidae
The mermithids represent a family of nematodes
that have become parasitic on other invertebrates.
Over half of the described genera parasitize only
aquatic insects. Two genera, Aranimermis and
Pheromermis, enter the immature stages of aquatic insects which are later consumed by terrestrial arthropods. However, the free-living stages of these and all
other aquatic mermithids (postparasitic juvenile,
adults, eggs, and infective juveniles) are found in the
aquatic habitat. Only the developing second- and
third-stage juveniles parasitize the aquatic stages of insects and other invertebrates; the other stages do not
take nourishment. Eggs and infective stage juveniles
are microscopic and normally not taken in samples,
but the postparasitic juveniles and adults are easily
seen and collected. Since the keys are based on adult
characters, postparasitic juveniles should be held in
water until they molt to the adult stage. One important character in identifying mermithids is the number
of hypodermal cords present. These can be determined
by cutting thin cross sections with a razor blade by
hand, and examining them under the microscope. The
hypodermal cords protrude through the muscle fields
in dorsal, ventral, lateral, and often the subdorsal and
subventral regions. Only genera known to occur in
North America are included.
1a.
Adult cuticle with cross-fibers; thick, robust white nematodes usually found along the edges of bogs, springs, and streams (parasites
of wasps, ants, and horseflies) (Fig. 11b)......................................................................Pheromermis Poinar, Lane, and Thomas, 1976
1b.
Adult cuticle without cross-fibers, robust or slender, white, green, pink, brown, or yellow nematodes found in lake, stream, and
pond beds...........................................................................................................................................................................................2
2a.
With four cephalic papillae..................................................................Pseudomermis de Man, 1903 (syn. Tetramermis Steiner, 1925)
2b.
With six cephalic papillae...................................................................................................................................................................3
3a (2b).
With single or fused (rare) spicules .....................................................................................................................................................4
3b.
With paired, separate spicules ............................................................................................................................................................8
4a (3a).
Spicule medium to long, more than twice anal body width ................................................................................................................5
4b.
Spicule short, less than twice anal body width....................................................................................................................................6
5a (4a).
Mouth terminal; spicules J-shaped; vulval flap present; [parasites of midges (Chironomidae)].......Lanceimermis Artyukhovsky, 1969
5b.
Mouth normally shifted ventrally; spicule curved but not J-shaped; vulva flap absent; [parasites of blackflies (Simuliidae), midges
(Chironomidae), and mayflies (Ephemeroptera)]................................................................................Gastromermis Micoletzky, 1923
6a(4b).
Spicule shorter than anal body width; amphids small; [parasites of mosquitoes (Culicidae)] ....................Perutilimermis Nickle, 1972
9. Nematoda and Nematomorpha
FIGURE 11 (a) A postparasitic juvenile aquatic mermithid emerging from an adult caddis
fly, Trichoptera (collected by H. Wolda and submitted by R. Schuster). (b) Adults of
Pheromermis pachysoma (Mermithidae) in a California spring bed. These nematodes have
emerged from parasitized queen wasps visiting the spring and will undergo molting, mating
and oviposition in the spring. (c) Various eggs and juvenile stages of freshwater nematodes
developing in a decaying, submerged leaf.
279
280
George O. Poinar, Jr.
6b.
Spicule longer than anal body width; amphids medium to large ........................................................................................................7
7a(6b).
Tail pointed; eight hypodermal cords; [parasites of midges (Chironomidae) and mosquitoes (Culicidae)] ...Hydromermis Corti, 1902
7b.
Tail rounded; six hypodermal cords; [parasites of midges (Chironomidae) and blackflies (Simuliidae)] .....Limnomermis Daday, 1911
8a (3b).
Vagina straight or nearly so, barrel- or pear-shaped ...........................................................................................................................9
8b.
Vagina S-shaped, U-shaped, or elongate with both ends curved........................................................................................................13
9a (8a).
Spicules shorter than cloacal body width..........................................................................................................................................10
9b.
Spicules equal to or longer than cloacal body width .........................................................................................................................11
10a (9a).
Head expanded into a bulblike shape with very thick cuticle; male lacking lateral – dorsal genital papillae; [parasites of midges
(Chironomidae)] .....................................................................................................................................Capitomermis Rubtsov, 1968
10b.
Head not expanded into a bulblike shape with thick cuticle; male with genital papillae on lateral-dorsal surface; [parasites of biting
midges (Ceratopogonidae)] ...................................................................................................................Heleidomermis Rubtsov, 1970
11a(9b).
Six hypodermal cords; [parasites of blackflies (Simuliidae)] .........................................................................Mesomermis Daday, 1911
11b.
Eight hypodermal cords ...................................................................................................................................................................12
12a(11b).
Spicules 2 – 4 times anal body width; [parasites of mosquitoes (Culicidae)] ............................................Romanomermis Coman,1961
12b.
Spicules 1-2 times anal body width; [parasites of mosquitoes (Culicidae) and midges (Chironomidae)] ...............................................
Octomyomermis Johnson, 1963
13a (8b).
Eight hypodermal cords ...................................................................................................................................................................16
13b.
Six hypodermal cords .......................................................................................................................................................................14
14a(13b).
Spicule length less than two times anal diameter; [parasites of midges (Chironomidae) and mosquitoes (Culicidae)] ...........................
Strelkovimermis Rubtsov, 1969
14b.
Spicule length three or more times cloacal diameter .........................................................................................................................15
15a(14b).
Spicules ten or more times body width at cloaca; vagina elongate with bends at both ends; postparasitic juveniles with distinct tail
appendage;[parasites of diving beetles (Dytiscidae)]................................................................Drilomermis Poinar and Petersen, 1978
15b.
Spicules 3 – 10 times body width at cloaca; vagina elongate with 3 – 6 irregular bends; postparasitic juveniles with indistinct or no
tail appendage; [parasites of spiders] ........................................................................................Aranimermis Poinar and Benton, 1986
16a(13a).
Vagina elongate, slightly curved; spicules shorter than anal body diameter; [parasites of mosquitoes (Culicidae)]................................
Culicimermis Rubtsov and Isaeva, 1975
16b.
Vagina S-shaped, ends distinctly curved; spicules range from shorter than anal body diameter to about 10 times tail diameter .......17
17a(16b).
Spicule length equal to or shorter than tail diameter; amphids small; cephalic crown well developed; [parasites of mosquitoes
(Culicidae)] ..............................................................................................................................................Empidomermis Poinar, 1977
17b.
Spicule length between one and ten times anal diameter; amphids medium – large, cephalic crown absent or only slightly developed;
[parasites of blackflies (Simuliidae)] ...............................................................................................................Isomermis Coman, 1953
NEMATOMORPHA
VII. INTRODUCTION
Members of the poorly known phylum Nematomorpha (Gordiacea) constitute a relict group that has
no clear or close relationship with any other living
forms. Fossil hairworms are rare and controversial.The
earlier determination of Gordius tenuifibrosus (Voigt,
1938) as a fossil hairworm (identified from a 15-mm
long fragment of subcuticular tissue in the Eocene
brown coals of the Geisel Valley near Halle, Germany)
is now considered questionable in light of similar
characters found on members of the Mermithidae
(Poinar, 1999). And while it has been proposed that
fossil palaeoscolecid worms are larval hairworms
(Xianguang and Bergstrom, 1994), similarities between
these two taxa are only superifical and provide no
evidence for phylogenetic relationship (Poinar, 1999).
The only unequivocal record of fossil Nematomorpha
are two hairworms, Paleochordodes protus Poinar
(1999), emerging from a cockroach in Dominican amber dated at 15 – 45 million years (Fig 20a). However,
the Nematomorpha certainly originated considerably
earlier (probably in the Early Paleozoic) as an offshoot
of now extinct lines. Chitwood (1950) considered that
Nematomorpha had the closest ties (albeit distant) with
nematodes and rotifers (the Acanthocephala and
9. Nematoda and Nematomorpha
Echinodermata were tied for the next closest kin). Using cladistic methods, Schmidt-Rhaesa (1998b) also
concluded that nematomorphs are closest to nematodes; however, it is difficult to establish lineages when
such basic characters as the number of molts during the
281
growth phase has not been confirmed. The body plan
of the Nematomorpha is unique in the animal kingdom
and even the sperm are considered aberrant and support the contention that this group separated quite
early from the other members of the Aschelminthes and
FIGURE 12 (a) Oldest known undisputed fossil nematode, Cretacimermis libani (Poinar
Acra and Acra) from 120 – 135 million-year-old Lower Cretaceous Lebanese amber. This
mermithid infected and developed in the aquatic larval stage of a midge (Chironomidae) and
was carried over into the adult host. (b) Two mermithid nematodes, Heydenius dominicanus
Poinar, in 20 – 40 million-year-old Dominican amber. These nematodes probably emerged
from an adult mosquito also enclosed in the amber (figure shows an adjacent cranefly adult
in the amber; the mosquito could not be included in the photo at this magnification).
282
George O. Poinar, Jr.
evolved independently thereafter (Lora Lami Donin
and Cotelli, 1977) (Fig. 15).
Some myths surround this group, perhaps associated with the proverbial “Gordian knot” (representing
a problem solvable only by drastic action as demonstrated by Alexander the Great cutting the knot that
could not be untied, which bound a chariot to a pole at
Gordium, the capital of Phrygia, in 333 BC). Another
myth, still believed by some today, is suggested by the
common name “hairworms” or “horsehair worms” indicating that they were supposed to have arisen from
horse hairs that fell into water. This belief was scientifically disproven by Leidy in 1870, when he observed
horse hairs placed in water over a period of many
months without “ . . . having had the opportunity of
seeing their vivification.”
Adult hairworms have been associated with the digestive and urogenital tract of humans and larval hairworms will burrow into a wide range of invertebrate
and vertebrate tissue, including human facial tissue
sometimes resulting in orbital tumors (Watson, 1960).
A review of the public health aspects of hairworms has
been provided by Cappucci (1982).
VIII. MORPHOLOGY AND PHYSIOLOGY
It is the free-living adults of hairworms that are
normally described because these stages are most
frequently encountered in sampling. They are dark
FIGURE 13
(rarely white), slender worms ranging in length from
several centimeters to 1 m and in width from 0.25 to
3 mm (Fig. 13a). The color of most adults varies from
yellowish to black; and although the anterior end is
generally attenuated, both tips tend to be obtusely
rounded or blunt (divided in some males and females). Because the cuticle is normally opaque, it is
impossible to examine the internal organs through the
body wall. Ultrastructural studies of a North American Gordius sp. revealed some interesting morphological features (Eakin and Brandenburger, 1974). The
cuticular structures that are such important taxonomic characters are actually sculpturing on the surface of the thin, superficial epicuticle (Figs. 13b, 16a,
19). This epicuticle is normally criss-crossed by
grooves or furrows, leaving small elevations of irregular areas (areoles) between them. The surfaces of the
areoles may be smooth (Fig. 19a) or may bear setae
(bristles) (Fig. 19b) or cuticular projections (tubercles)
(Fig. 19c,d), arranged singly or in clusters. These bristles and tubercles may also occur in the interareolar
furrows. Additional observations on the cuticular
structure of hairworms include investigations of
Paragordius varius by Zapotosky (1971), Parachordodes spp. by Smith (1991), G. aquaticus difficilis by
Smith (1994) and Chordodes morgani by Chandler
and Wells (1989). Many of these studies involve the
use of the scanning electron microscope which is a
useful tool for examining details of the surface sculpturing on the epicuticle.
(a) A hairworm (Nematomorpha) emerging from its orthopteran host. (b) Transverse section of
Gordius sp. C, cuticle; E, epidermis; EC, epicuticle; G, gut; M, muscles; ME, mesenchyme; NC, nerve cord; P,
pseudocoel; VNL, ventral neural lamella. [Courtesy of R. M. Eakin and J. L. Brandenburger.]
9. Nematoda and Nematomorpha
FIGURE 14 (a) Layer of eggs of Euchordodes nigromaculatus
Poinar scraped off a boulder in Cave Stream, Craigieburn forest,
New Zealand. Adhering the eggs to submerged substrates is an adaptation of hairworms living in fast flowing streams. (b) Ventral view of
tail of Gordius dimorphus Poinar showing bilobed tail with lobes
longer than wide, spherical cloacal opening located ventrally and a
semicircular postcloacal fold (arrow).
The cuticle, beneath the epicuticle, is composed of
many crossing layers of cylindrical, nonperiodic
collagenous fibers, spirally coiled along the length of the
adult (Fig. 16). Beneath the cuticle is the epidermal
layer, constructed of a single layer of interdigitated cells.
The musculature consists only of overlapping, longitudinal, flat cells; circular muscles are absent. Ultrastructural details of the musculature of G. aquaticus and
283
Nectomena munidae are provided by Schmidt-Rhaesa
(1998a). The pseudocoel is nearly filled with mesenchymal cells containing large clear vacuoles, which give the
tissue a foamy appearance. These cells are embedded in
a supporting collagenous matrix (Fig. 16b).
The mouth is located at the calotte or anterior tip
of the body (Fig. 18a). This region is often lighter in
color than the rest of the body and contains the nonfunctional pharynx, which, in turn, leads into a degenerate intestine lined with epithelium one cell in thickness. The gut epithelial cells of Gordius sp. contain
numerous microvilli that project into the lumen (Eakin
and Brandenburger, 1974).
In both sexes, the genital ducts empty into the intestine, forming a cloaca lined with cuticle (Fig. 18b).
Because adults do not feed, the cloaca is probably used
solely for reproductive purposes. All nematomorphs
are amphimictic. Males have paired cylindrical testes,
each of which connects with the cloaca via a separate
sperm duct. Females possess paired ovaries, and the
eggs, after passing through separate oviducts, enter the
cloaca independently.
Adults become sexually mature soon after emerging from their hosts and copulation occurs after the
male coils its posterior end around the terminus of the
female. The spermatozoa are either deposited as a spermatophore on the female terminus, from where they
migrate into the cloaca, or are deposited directly into
the female cloaca. Ultrastructural studies of the sperm
show that they are composed of a main cell body attached to a nuclear portion containing an elongated
nucleus with a helical configuration (Fig. 15a,b) (Lora
Lami Donin and Cotelli, 1977) (Schmidt-Rhaesa,
1997). The sperm structure is similar in many aspects
to that of mermithid nematodes (Poinar and HessPoinar, 1993). There are unique features, one of which
is what has been called a multivesicular complex, actually a number of flattened pockets, each filled with
mucopolysaccharide appearing substances which surround the nuclear portion of the spermatozoan (Fig.
15a,b). The eggs are deposited singly or in clusters
which often form elongate strings held together by secretions produced by the antrum (anterior portion of
the female cloaca). In certain species that occur in fast
flowing waters, ovipositional modifications occur.
While conducting studies on Euchordodes nigromaculatus in a stream in New Zealand, the present author
noted that the eggs were attached as a coating to rocks
and other submerged debris (Fig. 14a) (Poinar, 1989).
This unique manner of oviposition explains how
certain hairworms can maintain populations in fastflowing streams. Other investigators have noted eggs
of Euchordodes attached to twigs and other stationary
objects in the water (Bohall et al., 1997) (D.J.
Watermolen, personal observation).
284
George O. Poinar, Jr.
FIGURE 15 Electron micrographs of the lateral (a) and transverse (b) views of sperm cells of Neochordodes
occidentalis (Montgomery). M, main cell body; N, nucleus; G, glycogen deposit surrounding nucleus; O, membraneous organelles forming the multivesicular complex. (Photos courtesy of Roberta Poinar.)
The preparasitic stage that hatches from the egg is
morphologically quite different from the adult worm
and can, therefore, properly be called a larva. [Recall
that the term larva implies that some type of metamorphosis occurs before the adult stage.] The larva consists
of a presoma or preseptum (featuring an evaginable
proboscis armed with cuticular spines) and a body or
postseptum containing adult tissue primordia (Figs. 17,
18c,d). Ultrastructural studies of the larvae of
Paragordius varius demonstrated quite clearly the nature of the proboscis and spines (Zapotosky, 1974,
1975). It is unfortunate that so few larvae of described
nematomorph species are known, since they possess
their own specific characters which could be of taxonomic value, as noted by Poinar and Doelman (1974)
and Bohall et al. (1997).
9. Nematoda and Nematomorpha
FIGURE 16 (a) Surface view of the areolar pattern on the epicuticle
of Neochordodes occidentalis (Montgomery). (b) Raised areoles and
intra-areolae furrows on the epicuticle of a Gordius sp. DG, deep
groove separating areoles; SG, shallow grove separating areoles.
(Courtesy of R. M. Eaken and J. L. Brandenburger.) (c) Cross section
through the body wall of a Gordius sp. C, cuticle; E, epidermis; EC.
epicuticle; LF, cuticular fibers cut longitudinally; M, muscle plates;
XF, cuticular fibers cut crosswise. [Courtesy of R. M. Eaken and J. L.
Brandenbruger.]
FIGURE 17
285
(a) Figure of the preparasitic larva of Neochordodes
occidentalis (Montgomery). P, presoma (preseptum); B, body (postseptum); a, anus; g, globule. g,d, gland duct; i, intestine; i.g, intestinal
gland; m, mesenchyme cells; p.g, preintestinal gland; pr, protrusible
proboscis; r.m, retractor muscles; st, stylets; sp, circulet of spines; t,
tail spines (modified from Poinar and Doelman, 1974.). (b) Electron
micrograph of a larva of N. occidentalis encysted in the gut wall of a
tadpole of the tree frog Hyla regilla Baird and Girard. Note that the
surrounding cyst is composed of several layers. [Photo courtesy of
Roberta Poinar.]
FIGURE 18
(a) Anterior end of an adult N. occidentalis. Note the degenerate pharynx (arrow). (b) Ventral
view of male tail of N. occidentalis showing the elongate cloacal aperature (arrow). (c) Preparasitic larva of
N. occidentalis showing presoma (P), body (B), intestinal gland (C), and stylets (S). (d) Preparasitic larva of
N. occidentalis showing extruded secretions from the intestinal gland (arrows), stylet (ST), and spines (SP).
286
9. Nematoda and Nematomorpha
FIGURE 19
(a) Surface view of the body wall of Gordius dimorphus Poinar showing the absence of areoles
and the presence of parallel criss-cross fibers. (b) Surface view of the areoles of Euchordodes nigromaculatus
Poinar with minute spines present, especially in the furrows between the flattened areoles. (c) Surface view of
the areoles of Gordionus diblastus (Orley) showing variation in areole size and distribution. (d) Lateral view of
the raised areoles of G. diblastus showing size variability.
287
288
George O. Poinar, Jr.
IX. DEVELOPMENT AND LIFE HISTORY
The four stages in the life history of nematomorphs
are the egg, the preparasitic larva that hatches from the
egg, the parasitic larva that develops within an invertebrate, and the free-living, aquatic adult. Parasitic larvae
of all Nematomorpha mature only within invertebrates, which can be called the developmental or definitive hosts. A second type of host involved in the life
cycle of probably the great majority of hairworms is a
transport or paratenic host. The preparasitic larva enters the hemocoel and internal tissues of this host, but
does not develop further until the paratenic host is
eaten by a scavenger or predator. The paratenic host is
usually an invertebrate, but can also be a vertebrate
(e.g., tadpoles, fish, etc.). Poinar and Doelman (1974)
noted that when larvae of Neochordodes occidentalis
encysted in tadpoles (Hyla regilla), many of the cysts
were composed of several overlapping layers as is
shown in Figure 17b, in contrast to those normally
formed in insects. However, such encysted larvae of
N. occidentalis in the gut of tadpole paratenic hosts
showed no abnormal features.
Three types of life cycles have been described for
nematomorphs. The first is the direct type where the
egg hatches in water and the preparasitic larva is ingested by, and develops in, the definitive invertebrate
host. Such a cycle was demonstrated with Trichoptera
larvae (Stenophylax sp.) infected by Gordius sp.
(Dorier, 1930). The second type of cycle could be called
indirect-free-living, since after the preparasitic larva
hatches from the egg, it encysts on submerged leaves or
other detritus. As the water dries up (found in forms
inhabiting temporary ponds), the definitive host is infected by ingesting the cysts on the now exposed vegetation. Dorier (1930) indicated this type of development in a Gordius sp. that infected millipedes. For this
cycle to be effective, the cysts would have to be resistant to desiccation. Further confirmation of this type of
development is needed. The third type of cycle can be
called indirect-paratenic where, after hatching, the
preparasitic larva is ingested by an invertebrate or vertebrate. Hatching occurs when the mature preparasitic
larva forces itself through the wall of the egg (Fig.
20b). The parasite burrows into the tissues of this
paratenic host and then encysts, but does not develop
further. Only when the paratenic host is eaten by a
predator (carabid beetle, praying mantis, dragonfly) or
omnivore (various Orthoptera) does the parasite initiate development. This type of cycle occurs in the great
majority of investigated hairworms, including a Chordodes developing in praying mantids that ingest infected mayfly paratenic hosts (Inoue, 1962), a Gordius
developing in Dytiscus that feed on infected tadpole
(Rana temporaria) paratenic hosts (Blunck, 1922), Euchordodes nigromaculatus developing in stenopelmatids that fed on infected stoneflies, mayflies or caddisflies (Poinar, 1991), Gordius robustus developing in
crickets that were fed infected meal worms (Hanelt and
Janovy, 1999) and probably Neochordodes occidentalis
which readily infects and encysts in mosquitoes and
other aquatic insects (Poinar and Doelman, 1974). In
fact, it is apparent that this indirect-paratenic type of
cycle is the most typical type of development for hairwroms. The first two types of development mentioned
above need confirmation.
There is no evidence that the preparasitic larvae
can burrow directly through the outer body wall of either the paratenic or definitive host. In all cases, they
enter by way of mouth and encyst in the midgut (Fig.
20e) or burrow through the midgut and encyst in the
tissues of the body cavity. In cases of mass infection as
demonstrated in laboratory studies with larvae of N.
occidentalis invading mosquito larvae, the tissues may
be damaged to such a degree that the host dies (Poinar
and Doelman, 1974). In cases where the primary
paratenic host is eaten by an insect predator which is
not suitable for further parasitic development, the hairworm larvae may be able to re-encyst in the predator.
Encysted larvae in dobsonflies probably represented
cases of secondary paratenic hosts (Poinar, 1991).
Host immunity to hairworms has thus far been
documented only in the case of larvae entering or
remaining in paratenic hosts. The usual type of host reaction to such parasitic larvae is humoral melanization
which may occur immediately after entry as in the case
of N. occidentalis invading mosquito larvae (Fig.
20c,d) (Poinar and Doelman, 1974) or after the larvae
have encysted in the internal tissues of the paratenic
host as with hairworm larvae in caddis-flies, mayflies,
stoneflies (Fig. 20e) and especially chironomid larvae
(Poinar, 1991). Such melanization reactions often result
in the death of invading larvae (Fig. 20d), although in
cases where the host reaction occurred after encystment, it is possible that the parasites died of other
causes and were then melanized.
When parasitic development has been completed
and the hairworm is ready to emerge, the host must
come into contact with water. This poses no problem for
parasites in aquatic hosts, but can present obstacles
when the host is a terrestrial arthropod. Although entry
into water may occur accidentally, it is more likely that
the hosts suddenly have a desire to reach water. This
may result from partial desiccation due to the actions
of the parasite or be a consequence of some abnormal
stimuli from parasite products that affect physiological
centers of the host. When the host enters water, the
hairworms emerge and either slowly sink or initiate
FIGURE 20
(a) Two specimens of Paleochordodes protus Poinar (1999) that were in the process of emerging
from their cockroach host in 15 – 45 million-year-old Dominican amber. These represent the first unequivocal
fossil Nematomorpha. (b) Pre-parasitic larva of Neochordodes occidentalis ready to emerge from its egg. (c)
Host reaction showing initial stages of melanization of an invading larva of N. occidentalis that just penetrated
into the hemocoel of a fourth stage mosquito larva (experimental infection; see Poinar and Doelman, 1974).
(d) Melanized larva of N. occidentalis killed from the host reaction of a fourth stage mosquito larva (experimental infection; see Poinar and Doelman, 1974). (e) Dead, melanized encysted larvae of probably Euchordes
nigromaculatus in the intestinal wall of the stonefly, Stenoperla prasina (natural infection; see Poinar, 1991).
289
290
George O. Poinar, Jr.
TABLE II List of Host families Reported to contain the
Developing Stages of Nematomorphaa
Taxon
Phylum Annelida
Class Hirudinea
Order Rhynchobdellida
Family Glossiphoniidae
Order Pharyngobdellida
Family Erpobdellidae
Phylum Mollusca
Class Gastropoda
Subclass Pulmonata
Family Lymnaeidae
Phylum Arthropoda
Class Arachnida
Order Scorpionida
Family Vaejovidae
Order Ambylpygi
Family Phrynidae
Class Crustacea
Order Notostraca
Family Apodidae
Order Decapoda
Family Caridae
Class Chilopoda
Order Lithobiomorpha
Family Lithobiidae
Order Scolopendromorpha
Family Scolopendridae
Class Diplopoda
Order Glomerida
Family Glomeridae
Order Spirobolida
Family Spirobolidae
Order Julida
Family Julidae
Class Insecta
Order Odonata
Family Libellulidae
Order Orthoptera
Family Acrididae
Family Blattidae
Family Gryllacrididae
Family Gryllidae
Family Rhaphidophoridae
Family Stenopelmatidae
Family Tettigonidae
Order Mantodea
Family Mantidae
Order Phamsatida
Family Phasmatidae
Order Dermaptera
Family Forficulidae
Order Hemiptera
Family Notonectidae
Order Neuroptera
Family Sialidae
Order Lepidoptera
Family Saturnidae
Order Coleoptera
TABLE II
(Continued)
Taxon
Reference
Family Carabidae
Family Chrysomelidae
Family Dytiscidae
Family Silphidae
Family Tenebrionidae
Order Hymenoptera
Family Apidae
Blunck (1922)
Dorier (1930)
Blunck (1922)
Blunck (1922)
Baylis (1944)
Reference
Leidy (1878)
Sawyer (1971)
a
Chitwood and Chitwood (1937)
S. Williams (personal
communication)
Sciacchitano (1958)
Linstow (1878)
Linstow (1878)
Dorier (1930)
Dorier (1930)
Dorier (1930)
Cooper and Storck (1973)
Sahli (1972)
Heinze (1937)
Blunck (1922)
Sciacchitano(1958)
Poinar (1989)
Montogomery (1907)
Poulin (1996)
Poinar (1991)
Thorne (1940)
Sciacchitano (1958)
Römer (1895)
Sciacchitano (1958)
Cury (1946)
It is often impossible to determine whether authors are referring to
paratenic or developmental hosts in their reports. Questionable references have been omitted.
undulating body movements. Males are claimed to be
more active than females. As the parasites mature in the
host, they change from a light, cream color to a yellowish brown or dark brown color. At the time of emergence, most have already turned their natural color, but
in some cases, further darkening occurs after emergence.
Since there are so few records of known gordiids
from identified hosts, very little is known about host selection and specificity. Some hairworms are probably restricted developmentally to certain invertebrate genera,
while others may be able to develop in a wide range of
hosts. Most definitive hosts are medium-to-large bodied
predaceous or omnivorous arthropods. The great majority of freshwater nematomorphs have been collected
from representatives of the insect orders Coleoptera and
Orthoptera. Other host groups include myriapods
(Diplopoda and Chilopoda), crustaceans, and leeches. A
list of invertebrate families known to contain developing
stages of hairworms is presented in Table II. The diversity
of paratenic hosts is quite great, extending from trematodes (Cort, 1915) to vertebrates. Preparasites will attempt to burrow and encyst into the soft tissues of any
cold blooded organism that ingests them. The natural enemies of hairworms are poorly known. Fish ingest the
adults and the preparasites are probably fed upon by invertebrate micropredators. A Gordius adult that was
brought to nestlings by a parent bird (a tapaculo,
Scelorchilus rubecula) in a temperate rain forest in Chile
was submitted to the present author by Mary F. Willson
(personal correspondence, 1994), so birds can be listed as
hairworm predators. Predation by rock bass, brown
trout and crayfish has also been documented (Cochran
et. al., 1999).
Zalmon (1977)
Mellanby (1951)
X. SAMPLING
Camerano (1897)
Except for the marine genus Nectonema, all known
representatives of the Nematomorpha occur in freshwater. Their habitats range from watering troughs and
(Continues)
9. Nematoda and Nematomorpha
puddles to rivers, lakes and subterranean streams. Several
researchers have commented on their occurrence in the
Great Lakes (Watermolen and Haen, 1994; Cochran
et al., 1990). No special sampling technique has been developed for hairworms. They are large enough to be seen
with the naked eye and can be netted or lifted out of the
water by hand. Stream forms often accumulate in slower
moving water off to the side of the main current. Holes
dug into small streamlets will often catch and hold adults
that are being carried downstream and nets can be
placed across small streams to collect the unattached
adults. Late summer and spring are the best times to look
for the free-living adults. Although the long, whitish egg
strands of some species can sometimes be spotted in water, the preinfectives are too small to be found and are
usually noted only by chance when examining the
paratenic hosts. Perhaps dissecting paratenic hosts and
searching for encysted larvae is the easiest way to determine if hairworms are present in an aquatic system. The
eggs of those species adapted to fast-flowing streams can
be noted stuck to the surfaces of rocks and other debris.
Growing larvae in developmental hosts can be found
throughout the year, but are much less frequent than are
encysted larvae in paratenic hosts and are nearly impossible to keep alive if they are removed from their host prior
to completion of their development. No methods are
available for in vitro development or culture of nematomorphs. May (1919) was able to monitor development
by injecting preparasitic larvae of Gordius into the body
cavity of long-horned grasshopper hosts (Tettigonidae).
However, because of the fragile character of the preparasitic forms and the relatively long period of parasitic development (several months) coupled with the problem of
maintaining the hosts, few workers have even bothered
to culture hairworms in vivo.
XI. IDENTIFICATION
Adult hairworms can be preserved in 5‰ formalin
or 70% alcohol for identification purposes. All diagnostic characters are external features associated with
the head, tail, and epicuticular surfaces. The extremi-
291
ties can be removed and mounted in lactophenol or
glycerin after dehydration in an alcohol series. For cuticular examination, small slivers of epicuticle can be
removed from the body regions with a razor blade and
placed in lactophenol or dehydrated in glycerin. The
underlying epidermis and muscle tissue can then be
scraped away and the cuticular slice mounted (outer
surface up) in lactophenol or glycerin. This will expose
the areoles and their ornamentation. Some workers
prefer to clear the sections in xylene before mounting
them. The scanning electron microscope has been used
to reveal cuticular details and, although it can be a useful tool, workers should not base descriptions on characters only visible with the SEM since it is not practical
for many, including field biologists. Besides, adequate
diagnostic characters can normally be observed at least
with oil immersion objectives (1000 ). In a study
comparing the light and scanning electron microscope
in differentiating cuticular characters separating Parachordodes lineatus and P. violaceus, Smith (1991)
noted that while details were clearer with SEM, the
light microscope revealed the essential characters
needed to separate the species. Success in large part
with the light microscope depends greatly on skill in
preparing the sample. Both males and females should
be available for accurate identification; however, some
males can be identified alone using the following key.
The classification used here follows that presented
by Dorier (1965) in the Traité de Zoologie. It also incorporates the contributions of Heinze (1952) as well
as studies on North American forms by May (1919),
Leidy (1851), Carvalho (1942), Montgomery (1898a,
b, 1899, 1907), Poinar and Doelman (1974), Redlich
(1980), Chandler (1985), Chandler and Wells (1989),
Smith (1991; 1994), Pennak (1989) and others.
A. Taxonomic Key to Extant Genera of Nematomorpha
Although some of the genera included have not been
reported from North America, it is highly probable that
they as well as other still undescribed genera occur in the
Nearctic. Those genera with reported North American
representatives are marked with an asterisk.
1a.
Adults with natatory hairs; marine forms ........................................................................................Nectonematoidea (Rauther, 1930)
....................................................................................................................................................................*Nectonema Verrill, 1879
1b.
Adults lacking natatory hairs; freshwater forms ......................................................................................Gordioidea Rauther, 1930 2
2a (1b).
Cuticle covered with ridges; male cloaca terminal .....................................................................................Chordodiolus Heinze, 1935
2b.
Cuticle not covered with ridges, male cloaca subterminal...................................................................................................................3
3a (2b).
Cuticle smooth, lacking aeroles or with flat, smooth areoles; male tail bilobed (Fig. 14b), with a postcloacal fold (Fig. 14b); female
tail entire ......................................................................................................................................................Gordiidae May, 1919 4
3b.
Cuticle with distinct areoles usually ornamented with bristles or tubercles; male tail entire or if bilobed, then without a postcloacal
cresent; female tail entire or trilobed ........................................................................................................Chordodidae May, 1919 5
292
George O. Poinar, Jr.
4a (3a).
Lobes of male tail rounded (Fig. 14b); head region not attenuated ...............................................................*Gordius Linneaus, 1766
4b.
Lobes of male tail pointed; head region attenuated....................................................................................Acutogordius Heinze, 1952
5a (3b).
Male tail entire; female tail entire .......................................................................................................................................................6
5b.
Male tail bilobed; female tail entire or trilobed ..................................................................................................................................9
6a (5a).
Epicuticle with one type of areole, containing short bristles and tubercles * Neochordodes Carvallo, 1942 ........................................
6b.
Epicuticle with two or more types of areoles ......................................................................................................................................7
7a (6b).
Areoles containing prominent tubercles, crowns or papillae; female cloaca terminal ................................* Chordodes Creplin, 1847
7b.
Areoles without prominent tubercles, crowns or papillae; female cloaca subterminal........................................................................ 8
8a (7b).
Minute spines present on cuticle, especially in furrows between areoles (Fig. 19b); slender, small forms..........................Euchordodes
Heinze, 1937
8b.
Interareolar furrows without spines; normal-sized forms ...............................................................*Pseudochordodes Carvalho, 1942
9a (5b).
Female tail trilobed .............................................................................................................................* Paragordius Camerano, 1897
9b.
Female tail entire ..............................................................................................................................................................................10
10a (9b).
Areoles arranged in rows separated by ridges ............................................................................................Beatogordius Heinze, 1934
10b.
Areoles randomly arranged ..............................................................................................................................................................11
11a (10b). Interareolar furrows large, forming a network around the areoles...........................................................Semigordionus Heinze, 1952
11b.
Interareolar furrows normal .............................................................................................................................................................12
12a(11b).
A single type of areole present although they may vary in size and be single, bifid or trifid (Fig. 19c,d)...............................Gordionus
G. W. Muller, 1927
12b.
Two different types of areoles present ..............................................................................................................................................13
13a(12b).
Lobes on male tail longer than wide; adult head clearly flattened; pore canals present on larger areoles in males.................................
*Parachordodes Camerano, 1897
13b.
Lobes on male tail approximately as long as wide; adult head slightly flattened; pore canals absent or if present, then only in interareolar areas in males ..............................................................................................................................Paragordionus Heinze, 1935
LITERATURE CITED
Ayoub, S. M. 1977. Plant Nematology. An agricultural training aid. California Department of Food and Agriculture, Sacramento. 157 pp.
Baylis, H. A. 1944. Notes on the distribution of hairworms (Nematomorpha: Gordiidae) in the British Isles. Proceedings of the
Zoological Society of London 113: 193 – 197.
Blunck, H. 1922. Die Lebensgeschichte der im Gelbrand
schmarotzenden
Saitenewürmer.
Zoologischer
Anzeiger
54:111 – 132, 145 – 162.
Bohall, P. J., Wells, M. R., Chandler, C. M. 1997. External morphology of larvae of Chordodes morgani (Nematomorpha). Invertebrate Biology 116:26 – 29.
Bretschko, G. 1973. Benthos production of a highmountain lake:
Nematoda. Verhandlungen des Instituts für Vereinforschung in
Limnologica 18:1421 – 1428.
Camerano, L. 1897. Monografia dei Gordii. Memorie della Reale
Accademia delle Scienze di Torino 47:339 – 419.
Cappucci, Jr., D. T. 1982. Gordian worms (Hairworms): biological
and public health aspects, in: Steele, J. H. Editor-in-Chief, CRC
Handbook Series in Zoonoses, Section C: Parasitic Zoonoses,
Vol. II, CRC Press, Inc., Boca Raton, FL, pp. 193 – 203.
Carvalho, J. M. C. 1942. Studies on some Gordiaceae of North and
South America. Journal of Parasitology 28:213 – 222.
Chandler, C. M. 1985. Horsehair worms (Nematomorpha, Gordioidea) from Tennessee, with a review of taxonomy and distribution in the United States. Journal of the Tennessee Academy
of Science 60:59 – 62.
Chandler, C. M., Wells, M. R. 1989. Cutlcular features of Chordodes
morgani (Nematomorpha) using scanning electron microscopy.
Transactions of the American microscopy Society 108:
152 – 158.
Chitwood, B. G. 1950. Nemic relationships, in: Chitwood, B. G.
Chitwood, M. B., Eds., Introduction to Nematology. University
Park Press, Baltimore, MD, pp. 227 – 241.
Chitwood, B. G., Allen, M. W. 1959. Nemata, in: Edmondson, W. T.,
Ed., Fresh-water biology. 2nd ed., Wiley, New York, pp.
368 – 401.
Chitwood, B. G., Chitwood, M. B. 1937. Snails as hosts and carriers of nematodes and Nematomorpha. The Nautilus 50:
130 – 135.
Cobb, N. A. 1914. North American fresh-water nematodes. Transactions of the American Microscopical Society 33:35 – 100.
Cochran, P. A., Watermolen, D. J., Haen, G. L. 1990. Horsehair
worms (Phylum Nematomorpha) in Lake Michigan. Journal of
Great Lakes Research 16:485 – 487.
Cochran, P. A., Kinziger, A. P., Poly, W. J. 1999. Predation on Horsehair worms (Phylum Nematomorpha). Journal of Freshwater
Ecology 14:211–218.
Coe, W. R. 1937. Methods for the laboratory culture of nemerteans,
in: Needham, J. G., Ed., Culture methods for invertebrate animals. Dover, New York, pp. 162 – 165.
Cooper, C. L., Storck, T. W. 1973. Gordius sp., a new host record.
Ohio Journal of Science 73:228.
Corbel, J.-C. 1967. Les parasites des Orthopteres. Annales de Biologie 6:391 – 426.
9. Nematoda and Nematomorpha
Cort, W. W. 1915. Gordius larvae parasitic in a trematode. Journal
of Parasitology 1:198 – 199.
Cury, R. 1946. Molestias das abelhas. Biologico 12:241 – 254.
Dorier, A. 1930. Recherches biologiques et systematiques sur les Gordiaces. Travaux du laboratoire d’hydrobiologie et de pisciculture de l’ Universite de Grenoble, 22 année:1 – 183.
Dorier, A. 1965. Classe Gordiaces, in: Grasse, P.-P., Ed., Traite de
Zoologie. Vol. 4. Masson, Paris, pp. 201 – 1222.
Downing, J. A., Rigler, F. H., Eds. 1984. A manual on methods for
the assessment of secondary production in freshwaters. Blackwell, Oxford.
Eakin, R. M., Brandenburger, J. L. 1974. Ultrastructural features of a
Gordian worm (Nematomorpha). Journal of Ultrastructure Research 46:351 – 374.
Esser, R. P., Buckingham, G. R 1987. Genera and species of free-living
nematodes occupying fresh-water habitats in North America, in:
Veech, J. A., Dickson, D. W. Eds. Vistas on Nematology. Society
of Nematologists, Hyaltsville, MD, pp. 477 – 487.
Esser, R. P., Buckingham, G. R., Bennett, C. A., Harkcom, K. J.
1985. A survey of phytoparasitic and free-living nematodes
associated with aquatic macrophytes in Florida. Proceedings of
the Florida Soil and Crop Science Society 44:150 – 155.
Ferris, V. R., Ferris, J. M. 1972. Nematode community structure: a
tool for evaluating water resource environments. Technical report No. 30. Water Resources Research Center, Purdue University, Lafayette, IN.
Ferris, V. R., Ferris, J. M., Tjepkema, J. P. 1973. Genera of freshwater nematodes (Nematode) of Eastern North America. U.S.
Environmental Protection Agency Identification Manual No.
10. 38 pp.
Filipjev, I. N., Schuurmans Stekhoven, Jr., J. H. 1959. Agricultural
Helminthology. Brill, Leiden, The Netherlands.
Flint, R. W. 1976. The natural history, ecology and proauction of the
crayfish, Pacifastacus leniusculus, in a subalpine lacustrine environment. Ph.D. Thesis, University of California, Davis.
Gerber, K., Smart, Jr., G. C. 1987. Plant-parasitic nematodes associated with aquatic vascular plants, in: Veech, J. A., Dickson,
D. W., Eds. Vistas on Nematology. Society of Nematologists,
Hyaltsville, MD, pp. 488 – 501.
Goodey, J. B. 1963. Soil and Freshwater Nematodes. Methuen, London.
Hanelt, B. Janovy, Jr., J. 1999. The life cycle of a horsehair worm,
Gordius robustus (Nematomorpha: Gordioidea). Journal of
Parasitology 85:139 – 141.
Hantzschel, W. 1975. Trace fossils and problematics, in: Moore,
R. C., Ed., Treatise on Invertebrate Paleontology. Part W,
Supplement 1. The Geology Society of America. Boulder, CO,
pp. 1 – 269 .
Heinze, K. 1937. Die Saitenwürmer (Gordioidea) Deutschlands. Eine
systematisch-faunistische Studie über insektenparasiten aus der
Gruppe der Nematomorpha. Zeitschrift für Parasitenkunde
9:263 – 344.
Heinze, K. 1952. Über Gordioidea, eine systematische Studie über Insektenparasiten aus der Gruppe der Nematomorpha. Zeitschrift
für Parasitenkunde 7:657 – 678.
Heyns, J., Coomans, A. 1980. Freshwater nematodes from South
Africa. 5. Chronogaster Cobb, 1913. Nematologica 26:
245 – 265.
Hoeppli, R. J. C. 1926. Studies of free-living nematodes from the
thermal waters of Yellowstone Park. Transactions of the American Microscopical Society 45:234 – 255.
Holme, N. A., McIntyre, A. O., Eds. 1971. Methods for the study of
marine benthos. IBP Handbook No. 16. Blackwell, Oxford.
Holopainen, I. J., Paasivirta, L. 1977. Abundance and biomass of the
meozoobenthos in the oligotrophic and mesohumic lake Paa-
293
jarvi, southern Finland. Annales Zoologica Fennland
14:124 – 134.
Hulings, N. C., Gray, J. S., Eds. 1971. A manual for the study of
meiofauna. Smithsonian Contributions to Zoology 78:84 pp.
Hummon, W. D. 1981. Extraction by sieving: a biased procedure in
studies of stream meiofauna. Transactions of the American Microscopical Society 100:278 – 284.
Inoue, I. 1962. Studies on the life history of Chordodes japonensis, a
species of Gordiaceae. III. The mode of infection. Annotationes
Zoological Japonenses 35:12 – 19.
Issel, R. 1906. Sulla termobiosi negli animal) acquatici. Ricerche faunistiche e biologiche. Attidella Societa Ligustica di Scienze natural) e geografiche 17:3 – 72.
Jacobs, L. J. 1984. The free-living inland aquatic nematodes of
Africa — a review. Hydrobiologia 113:259 – 291.
Kirjanova, E. S. 1958. On the structure of the copulative organs of
males of the freshwater hairworms (Nematomorpha, Gordioidea). Zoological Zhurnal 37:359 – 372. (In Russian.)
Leidy, J. 1870. The gordius, or hairworm. The American Entomologist and Botanist 2:193 – 197.
Leidy, J. 1878. On Gordius infesting the cockroach and leech. Proceedings of the Academy of Natural Sciences of Philadelphia
30:383.
Linstow, O. von. 1878. Compendium der Helminthologie, Hanover.
Lora Lami Donin, C., Cotelli, F. 1977. The rod-shaped sperm of
Gordioidea (Aschelminthes, Nematomorpha). Journal of Ultrastructural Research 61:193 – 200.
May, H. G. 1919. Contributions to the life histories of Gordius robustus Leidy and Paragordius varius (Leidy). Illinois Biological
Monographs 5:1 – 119.
Mellanby, H. 1951. Animal life in freshwater. A guide to freshwater
invertebrates. 4th ed., Methuen, London.
Meyl, A. H. 1954. Beiträge zur Kenntnis der Nematodenfauna vulkanisch erhitzter Biotope. Zeitschrift für Morphologie und Ökologie der Tiere 42:421 – 448.
Micoletzky, H. 1922. Die freilebenden Erd-Nematoden. Archiv für
Naturgeschichte A87: 1 – 650.
Montgomery, Jr., T. H. 1907. The distribution of the North American Gordiacea, with descriptions of a new species. Proceedings
of the Academy of Natural Sciences of Philadelphia, 59:
270 – 272.
Montgomery, Jr., T. H. 1899. Synopses of North American invertebrates. II. Gordiaces (hairworms). American Naturalist 33:
647 – 652.
Montgomery, Jr., T. H. 1898a. The Gordiacea of certain American
collections with particular reference to the North American
fauna. Bulletin of the Museum of Comparative Zoology 32:
1 – 59.
Montgomery, Jr., T. H. 1898b. The Gordiacea of certain American
collections, with particular reference to the North American
fauna. II. Proceedings of the California Academy of Science
1:333 – 344.
Moussa, M. T. 1969. Nematode fossil tracks of Eocene age from
Utah. Nematologica 15:376 – 380.
Nalepa, T. F., Quigley, M. A. 1983. Abundance and biomass of the
meiobenthos in nearshore Lake Michigan with comparisons to
the macrobenthos. Journal of Great Lakes Research 9:530 – 547.
Nicholas, W. L. 1984. The biology of free-living nematodes. Clarendon Press, Oxford.
Nichols, J. A. 1979. A simple flotation technique for separating
meiobenthic nematodes from fine-grained sediments. Transactions of the American Microscopical Society 98:127 – 130.
Nuttycombe, J. W. 1937. Wheat-grain infusion, in: Needham, J. G.
Ed. Culture methods for invertebrate animals. Dover, New
York, pp. 135 – 136.
294
George O. Poinar, Jr.
Oden, B. J. 1979. The freshwater littoral meiofauna in a South Carolina reservoir thermal effluents. Freshwater Biology 9:291 – 304.
Pennak, R. W. 1978. Fresh-water invertebrates of the United States.
2nd ed., Wiley, New York.
Pennak, R. W. 1989. Freshwater invertebrates of the United States.
3rd ed., Wiley, New York.
Petersen, J. J. 1972. Procedures for the mass rearing of a mermithid
parasite of mosquitoes. Mosquito News 32:226 – 230.
Poinar, Jr., G. O. 1983. The Natural History of Nematodes. PrenticeHall. Englewood Cliffs, N J.
Poinar, Jr., G. O. 1984. Heydenius dominicanus n. sp. (Nematoda:
Mermithidae), a fossil parasite from the Dominican Republic.
Journal of Nematology 16:371 – 375.
Poinar, Jr., G. O. 1989. Unpublished observations in Australia and
New Zealand.
Poinar, Jr., G. O. 1991. Hairworm (Nematomorpha: Gordioidea)
parasites of New Zealand wetas (Orthoptera: Stenopelmatidae)
Canadian Journal of Zoology 69:1592 – 1599.
Poinar, Jr., G. O. 1999. Paleochordodes protus n.g., n.sp. (Chordodidae:Nematomorpha), parasites of a fossil cockroach, with a
critical examination of other fossil hairworms and helminthes
of extant cockroaches (Blattaria: Insecta). Invertebate Biology
(118:109–115.).
Poinar, Jr., G. O., Doelman, J. J. 1974. A reexamination of Neochordodes occidentalis (Mont.) comb. n. (Chordodidae: Gordioidea):
larval penetration and defense reaction in Culex pipiens L. Journal of Parasitology 60:327 – 335.
Poinar, Jr., G. O., Hansen, E. 1983. Sex and reproductive modifications
in nematodes. Helminthological Abstract, Series B 52:145–163.
Poinar, Jr., G. O., Hess, R. 1988. Protozoan diseases of nematodes,
in: Poinar, G. O. Jr. Jansson, H.-B., Eds., Diseases of Nematodes. Vol. 1, pp. 103 – 131, CRC Press, Boca Raton, FL.
Poinar, Jr., G. O., Hess-Poinar, R. 1993. The fine structure of Gastromermis sp. (Nematoda: Mermithidae) sperm. Journal of Submicroscopic Cytology and Pathology 25:417 – 431.
Poinar, Jr., G. O., Sarbu, S.M. 1994. Chronogaster troglodytes sp.n.
(Nemata: Chronogasteridae) from Movile Cave, with a review
of carvernicolous nematodes. Fundamental and applied Nematology 17:231 – 237.
Poinar, Jr., G.O., Acra, A., Acra, F. 1994. Earliest fossil neam
tode (Mermithidae) in Cretaceous Lebanese amber. Fundamental and applied Nematology 17:475 – 477.
Por, F. D., Moary, D. 1968. Survival of a nematode and an
oligochaete species in the anaeorbic benthal of Lake Tiberios,
Oikos 19:388 – 391.
Poulin, R. 1996. Observations on the free-living adult stage of
Gordius dimorphus (Nematomorpha: Gordioidea), Journal of
Parasitology 82:845 – 846.
Rahm, G. 1937. Grenzen des Lebens? Studien in heissen Quellen.
Forschungen und Fortschritte 13:381 – 387.
Redlich, A. 1980. Description of Gordius attoni sp.n. (Nematomorpha, Gordiidae) from Northern Canada. Canadian Journal of
Zoology 58:382 – 385.
Riemann, F., Schrage, M. 1978. The mucus-trap hypothesis on feeding of aquatic nematodes and implications of biodegradation
and sediment texture. Oecologia 34:75 – 88.
Römer, F. 1895. Die Gordiiden des Naturhistorischen Museums in
Hamburg. Zoologische Jahrbücher, Abtheilung fur Systematik,
Geographie und Biologie der Thiere 8:790 – 803.
Sahli, F. 1972. Modifications des caracteres sexuel secondaires males
chez les Julidae (Myriapods, Diplopoda) sous l’influence de
Gordiaces parasites. Comptes Rendus de l’Academie des Sciences 274:900 – 903.
Sawyer, R. T. 1971. Erpobdellid leeches as new hosts for the Nematomorpha, Gordius. Journal of Parasitology 57:285.
Schiemer, F. 1978. Verteilung und Systematik der freilebenden Nematoden des Neusiedlersee. Hydrobiologie 58:167 – 194.
Schiemer, F. 1983. Comparative aspects of food dependence and energetics of freeliving nematodes. Oikos 41:32 – 42.
Schiemer, F. 1987. Nematoda, in: Pandian, T. J., Vernberg, F. J. Eds.,
Animal energetics. Vol. 1: Protozoa through Insecta. Academic
Press, New York, pp. 185 – 215.
Schmidt-Rhaesa, A. 1997. Ultrastructural observations of the male
reproductive system and spermatozoa of Gordius aquaticus L.
1758 (Nematomorpha). Invertebrate Reproduction and Development 32: 31 – 40.
Schmidt-Rhaesa, A. 1998a. Muscular ultrastructure in Nectonema
munidae and Gordius aquaticus (Nematomorpha). Invertebrate
Biology 117:38 – 45.
Schmidt-Rhaesa, A. 1998b. Phylogenetic relationships of the Nematomorpha- a discussion of current hypothesis. Zoologischer
Anzeiger 236:203 – 216.
Sciacchitano, I. 1958. Gordioidea del Congo Belga. Annales Museum
de la Republic de Congo Belge 67:7 – 111.
Smith, D. G. 1991. Observations on the morphology and taxonomy
of two Parachordodes species (Nematomorpha, Gordioidea,
Chordidae) in southern New England (USA). Journal of Zoology (London) 225: 469 – 480.
Smith, D. G. 1994. A reevaluation of Gordius aquaticus difficilis
Montgomery, 1898 (Nematomorpha, Gordioidea, Gordiidae).
Proceedings of the Academy of Natural Sciences of Philadelphia
145:29 – 34.
Southey, J. F. 1978. Laboratory methods for work with plant and soil
nematodes. Tech. Bull. No. 2. Ministry of Agriculture, Fisheries
and Food, London.
Stirewalt, M. A. 1937. The culture of Microstomum, in: Needham,
J. G., Ed., Culture methods for invertebrate animals. Dover,
New York, pp. 149 – 150.
Strayer, D. 1985. The benthic micrometazoans of Mirrow Lake, New
Hamphire. Archiv für Hydrobiologie, Supplement 72:287 – 426.
Thorne, G. 1940. The hairworm, Gordius robustus Leidy, as a parasite of the Mormon cricket, Anabrus simplex Haldeman. Journal of the Washington Academy of Science 30:219 – 231.
Tarjan, A. C., Esser, R. P., Chang, S. L. 1977. An illustrated key to
nematodes found in freshwater. Journal of Water Pollution Control Federation 49:23182 – 337.
Tsalolikhin, S. Y. 1983. The nematode families Tobrilidae and Tripylidae: World fauna. Nauka, Leningrad. (In Russian.)
Viglierchio, D. R., Schmidt, R. V. 1983. On the methodology of nematode extraction from field samples: Comparison of the methods for soil extraction. Journal of Nematology 15:450 – 454.
Voigt, E. 1938. Ein fossiler Saitenwurm (Gordius tenuifibrosus n.sp.)
aus der Eozänen Braunkohle des Geiseltales. Nova Acta
Leopoldina, new series 5:351 – 360.
Warwick, R. M., Price, R. 1979. Ecological and metabolic studies on
free-living nematodes from an estuarine mudflat. Estuary
Coastal Marine Science 9:257 – 271.
Watermolen, D. J., Haen, G. L. 1994. Horsehair worms (Phylum Nematomorpha) in Wisconsin, with notes on their occurrence in
the Great Lakes. Journal of Freshwater Ecology 9:7 – 11.
Watson, J. M. 1960. Medical Helminthology. Bailliere, London.
Winterbourn, M. J., Brown, T. J. 1967. Observations on the faunas
of two warm streams in the Taupo Thermal Region. New
Zealand Journal of marine and freshwater Resources 1:38 – 50.
Xianguang, H., Bergstrom, J. 1994. Palaeoscolecid worms may be
nematomorphs rather than annelids. Lethaia 27:11 – 17.
Zalmon, F. G. 1977. A nematomorph parasitizing a back swimmer
(Hemiptera: Notonectidae). The American Midland Naturalist
97:229 – 230.
Zapotosky, J. E. 1971. The cuticular ultrastructure of Paragordius
9. Nematoda and Nematomorpha
295
varius (Leidy, 1851) (Gordioidea, Chordodidae). Proceedings of
the Helminthological Society of Washington 38:228 – 236.
Zapotosky, J. E. 1974. Fine structure of the larval stage of
Paragordius varius (Leidy, 1851) (Gordioidea: Paragordiidae).
1. The Preseptum. Proceedings of the Helminthological Society
of Washington 41:209 – 221.
Zapotosky, J. E. 1975. Fine structure of the larval stage of
Paragordius varius (Leidy, 1851) (Gordioidea: Paragordiidae).
2. The postseptum. Proceedings of the Helminthological Society
of Washington 42:103 – 111.
Zullini, A. 1976. Nematodes as indicators of river pollution. Nematology Mediterranean 4:13 – 22.
APPENDIX 9.1 Systematic Arrangement of the
Nematode Generaa
APPENDIX 9.1 (Continued)
Phylum Nematoda
Class Adenophorea
Subclass Chromadorida
Order Araeolaimida
Suborder Araeolaimina
Superfamily Axonolaimoidea
Family Axonolaimidae (Cylindrolaimus)
Superfamily Leptolaimoidea
Family Bastianiidae (Bastiania)
Family Leptolaimidae (Rhabdolaimus)
Superfamily Camacolaimidae
Family Camacolaimidae (Aphanolaimus, Paraphanolaimus)
Superfamily Plectoidea
Family Chronogasteridae (Chronogaster)
Family Plectidae (Anonchus, Plectus)
Family Teratocephalidae (Euteratocephalus,
Teratocephalus)
Order Monhysterida
Superfamily Monhysteroidea
Family Monhysteridae (Monhystera, Monhystrella,
Prismatolaimus)
Order Chromadorida
Suborder Chromadorina
Superfamily Chromadoridea
Family Chromadoridae (Chromadorita, Punctodora)
Suborder Cyatholaimina
Superfamily Cyatholaimoidea
Family Cyatholaimidae (Achromadora, Ethmolaimus,
Odontolaimus, Paracyatholaimus, Prodesmodora
Family Microlaimidae (Microlaimus)
Order Enoplida
Suborder Enoplina
Superfamily Tripyloidea
Family Tripylidae (Tobrilus, Tripyla)
Family Alaimidae (Alaimus, Amphidelus)
Family Ironidae (Cryptonchus, Ironus)
Suborder Oncholaimina
Superfamily Oncholaimoidea
Family Oncholaimidae (Mononchulus, Oncholaimus)
Order Dorylaimida
Suborder Dorylaimina
Superfamily Dorylaimoidea
Family Dorylaimidae (Actinolaimus, Dorylaimus,
Eudorylaimus, Mesodorylaimus, Paractinolaimus)
Family Nygolaimidae (Nygolaimus)
Superfamily Leptonchoidea
(Continues)
Family Campydoridae (Aulolaimoides)
Order Mononchida
Suborder Mononchina
Superfamily Mononchoidea
Family Mononchidae (Anatonchus, Mononchus,
Prionchulus)
Order Mermithida
Suborder Mermithina
Superfamily Mermithoidea
Family Mermithidae (Aranimermis,
Capitomermis, Cretacimermis, Culicimermis,
Drilomermis, Empidomermis,
Gastromermis, Heleidomermis,
Heydenius, Hydromermis, Isomermis,
Lanceimermis, Limnomermis,
Mesomermis, Octomyomermis,
Perutilimermis, Pheromermis,
Pseudomermis, Romanomermis,
Strelkovimermis)
Class Secernentea
Order Tylenchida
Suborder Tylenchina
Superfamily Hoplolaimoidea
Family Hoplolaimidae (Hirschmaniella)
Superfamily Tylenchoidea
Family Tylenchidae (Tylenchus)
Superfamily Atylenchoidea
Family Atylenchidae (Atylenchus)
Suborder Criconematina
Superfamily Hemicyclcophoroidea
Family Hemicycliophoridae (Hemicycliophora)
Order Aphelenchida
Suborder Aphelenchina
Superfamily Aphelenchoidoidea
Family Aphelenchoididae (Aphelenchoides, Seinura)
Superfamily Aphelenchidae
Family Aphelenchidae (Aphelenchus)
Order Rhabditida
Suborder Diplogasteroidea
Superfamily Diplogasteroidea
Family Diplogasteridae (Rhabditolaimus)
Superfamily Rhabditoidea
Family Cephalobidae (Cephalobus, Eucephalobus)
Family Rhabditidae (Caenorhabditis, Mesorhabditis,
Pellioditis)
Family Daubayliidae (Daubaylia)a
a
The above include all genera cited in this chapter.
This Page Intentionally Left Blank
10
MOLLUSCA: GASTROPODA
Kenneth M. Brown
Department of Biological Sciences
Louisiana State University
Baton Rouge, Louisiana 70803
I. Introduction
II. Anatomy and Physiology
A. External and Internal
Morphology
B. Organ System Function
C. Environmental Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
I. INTRODUCTION
Gastropods are the most diverse class of the phylum Mollusca, with anywhere from 40,000 to 100,000
species, depending on the authority (Bieler, 1992; Ponder and Lindberg, 1997). In North America, there are
14 families, 88 genera and 659 species of freshwater
snails (Bogan, 1999). Snails are common organisms
along the margins of lakes and streams. They feed on
detritus, graze on the periphyton covering of macrophytes or cobble, or even float upside down at the water surface, supported by the surface tension, and feed
on algae trapped in the same fashion (Fig. 1). In fact,
gastropods often control the amount and composition
of periphyton in both lotic and lentic environments.
They are also the basis of food chains dominated by
sport fish. Predators, by controlling snail populations,
may indirectly facilitate algal producers. Freshwater
snails also have extensive intraspecific variation in life
histories, productivity, morphology and feeding habits
that adapt them to live in uncertain freshwater habitats. There is increasing concern, however, that many
riverine gastropod populations in the southeastern
United States are currently endangered because of
habitat alterations such as impoundments.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
D. Evolutionary Relationships
IV. Collecting and Culturing Freshwater
Gastropods
V. Identification of the Freshwater
Gastropods of North America
A. Taxonomic Key to Families and
Selected Genera of Freshwater
Gastropodas
Literature Cited
Snails are soft bodied, unsegmented animals, with
a body organized into a muscular foot, a head, a visceral mass containing most of the organ systems, and a
fleshy mantle which secretes the calcareous shell. Gastropods have a univalve shell and possess a filelike
radula used in feeding on the periphyton coverings of
rocks or plants. Traditionally, gastropods were divided
into three subclasses: Prosobranchia (including 53% of
modern species), Opistobranchia (4%), and Pulmonata
(43%). Prosobranch snails possess a gill (ctenidium)
and a horny (flexible) or calcareous operculum, or
“trap door,” which is pulled in after the foot to protect
the animal. Pulmonates have secondarily re-invaded
freshwaters from the terrestrial habitats used by their
ancestors, use a modified portion of the mantle cavity
as a lung, and lack an operculum.
Later authors divided the prosobranchs into the
ancestral archeogastropods, the mesogastropods, and
most-derived neogastropods, with a trend toward reduction in number of gills, auricles, and kidneys. Cox
(1960) combined the meso- and neogastropods into the
Caenogastropoda. More recent authors have also considered the valvatids to be more closely allied to the
opistobranchs and pulmonates than to the prosobranch
groups, and they have suggested combining the three
297
298
Kenneth M. Brown
II. ANATOMY AND PHYSIOLOGY
A. External and Internal Morphology
1. Shell
FIGURE 1 An adult Lymnaea stagnalis (length about 5 cm),
supported by the surface tension, is foraging upside down at the
water surface.
former groups into the Heterogastropoda (Kosuge,
1966). Cephalopods are thought to be the closest sister
group to the gastropods, and patellid limpets (a marine, intertidal group) are thought to be the most primitive gastropods (Ponder and Lindberg, 1997). More details on the time line of gastropod evolution are given
by Bandel (1997).
The first gastropods are thought to have had coiled
shells, and several trends have occurred throughout
gastropod evolution (Ponder and Lindberg, 1997). Torsion (a 180° rotation of the gut so that the anus and
mantle cavity lie anterior) is a functional homology of
all gastropods, although it is secondarily reduced in
opistobranchs and pulmonates. Anatomical complexity
has also become secondarily simplified, including the
radula, the circulatory system, and digestive anatomy.
External fertilization and pelagic larvae are also considered ancestral traits, with internal fertilization, direct
development and encapsulated eggs being derived
traits. In contrast, respiratory systems, neuro-secretory
structures, and life histories have increased in complexity through evolutionary time. A detailed introduction
to gastropod biology can be found in Fretter and Graham (1962), and in volumes on the general biology of
molluscs edited by Wilbur (1983) or the biology of pulmonates edited by Fretter and Peake (1975, 1978).
The structure of the shell is important in systematics. Shells can have a simple conical shape (Fig. 2A), as
in the limpets, family Ancylidae, with new shell material secreted at the margin. Second, the shell can be
planospiral (with the whorls all in one plane), as in the
pulmonate family Planorbidae (Fig. 2B). Whorls may
be elevated into a “spire,” as in the pulmonate families
Physidae and Lymnaeidae, and in various prosobranch
families (Fig. 2 C, D). Russell-Hunter (1983) discusses
how new shell material is secreted by the mantle for
each of these three basic shapes. If shells are placed
with the spire facing away from the observer and the
aperture (opening from which the foot extends) upward, shells with the aperture on the left are sinistral
(e.g., the physids), whereas those on the right are dextral (e.g., the lymnaeids and prosobranchs). Spiral
shells have a central supporting member, the columella,
similar to the center support of a spiral staircase. The
columella adds to the strength of the shell, and provides an attachment point for the soft parts via the columellar muscle.
A spiral shell (Fig. 2D) is convenient for illustrating
terminology used in systematics. The pointed end of
the shell above the aperture is the apex. Shell length in
spiral shells is measured from the apex to the lower tip
of the aperture, while the greatest diameter is used in
planospiral shells. The spire is separated into a number
FIGURE 2
Basic anatomy of the shell, including shell architecture
(A, conical; B, planospiral; C and D, spiral, with major shell features
indicated in D) and three types of opercula (E, concentric, F,
paucispiral, and G multispiral).
10. Mollusca: Gastropoda
of whorls by sutures (Fig. 2). The most apical whorl is
the nuclear whorl or protoconch, the initial shell of the
newly hatched snail or “spat.” The final and biggest
whorl (representing the most recent shell growth) is the
body whorl, ending in the aperture. Whorls may be
rounded and sutures deep and well defined (as in a typical lymnaeid shell, Fig. 2D), or they may be flattened
and sutures shallow, as in the prosobranch family Pleuroceridae (Fig. 2C). Spiral shells thus have a variety of
shapes. If the whorls are flat, the shell is termed coneshaped, while moderately inflated whorls produce a
“subglobose” shell, and shells that are almost circular
are globose; there is a continuum between these three
shapes. Shell thickness varies from thin and fragile (as
in many pulmonates) to thick and resistant to crushing
(as in the prosobranchs).
Part of the aperture is often reflected over the body
whorl at the columella (Fig. 2D) to form an inner lip. If
there is a channel between the inner lip and the body
whorl, the shell is umbilicate or perforate (the opening
of the channel is called the umbilicus and leads up and
inward into the columella). Imperforate shells lack an
umbilicus. While freshwater shells are not as ornate as
their marine relatives (Vermeij and Covich, 1978),
shells may have spines, ridges running along the margins of the whorls (called carina), colored bands, or
small malleations (hammerings) on the surface. Ridges
at right angles to the whorls are called costae, while
smaller ridges running spirally along the whorls are lirae. The operculum is useful in classifying prosobranchs. If the growth lines lie completely within each
other, the operculum is concentric; whereas, lines
arranged in a spiral are termed multispiral or paucispriral (see examples in Fig. 2 E – G). Further discussion on
shell sculpture is given in Fretter and Graham (1962)
and Burch (1989).
The shell is composed of an outer periostracum of
organic (mostly protein) composition which may limit
shell abrasion or dissolution of shell calcium carbonate
by acid waters. Beneath the periostracum is a thick
layer of crystalline calcium carbonate with some
protein material as a matrix for the calcium crystals
(Russell-Hunter, 1978). Calcium carbonate is either
absorbed directly from water or is sequestered from
food (McMahon, 1983). Snails in calcium-poor waters
(5 mg / L, Lodge et al., 1987) expend energy to absorb calcium against a gradient, and many species are
therefore limited to calcium-rich habitats.
2. Soft Anatomy
The soft parts are separated into: (1) head, (2)
foot, (3) visceral mass, and (4) the mantle (Fig. 3).
Aquatic pulmonates and prosobranchs possess eyes at
the base of their tentacles, unlike terrestrial pulmonates
299
FIGURE 3 Basic internal anatomy (with shell removed) of a
planorbid pulmonate (above, after Burch, 1989), and a pleurocerid
prosobranch (below, after Pechenik, 1985).
whose eyes are at the tips of the tentacles. The muscular foot is provided with both cilia and secretory
epithelium to secrete mucus for locomotion, as well as
pedal muscles which produce waves of contraction to
push the animal forward. The visceral hump includes
most of the organs of digestion and reproduction. The
mantle covers the visceral mass, and underlays the
shell, which it secretes. The anterior mantle, over the
head, possesses a mantle cavity, where the gill or ctenidium is located in prosobranchs. Gastropods have ganglia innervating each of these areas. Further information on internal anatomy can be found in Fretter and
Graham (1962), Barnes (1987), Pechenik (1985), or
Hyman (1967).
B. Organ System Function
1. Reproductive System
Prosobranchs are usually dioecious, and males use
the enlarged right tentacle as a copulatory organ (in the
viviparids), or possess a specialized penis or verge (in
the hydrobiids, pomatiopsids, and valvatids) or have
no copulatory organ (thiarids and pleurocerids). The
penis is usually located behind the male’s right tentacle.
Many pleurocerids lay clutches of a few eggs, whereas
300
Kenneth M. Brown
FIGURE 4
Anatomy of the reproductive system of the monoecious pulmonate Physella (after
Duncan, 1975). Sperm are produced in the ovotestis, stored in the seminal vesicle, exit via vas deferens, and are inserted into the vagina of a second individual via the introvertable penis; or sperm may
fertilize the same individual’s eggs in the hermaphroditic duct. Most foreign sperm are hydrolyzed in
Physa in the bursa copulatrix, but some do fertilize eggs in the fertilization pocket. The albumen
gland secretes yolk around the egg, and the oviduct wall secretes both the egg membrane and the
jelly and external wall of the egg case.
viviparids produce eggs which hatch and develop in
a fold of the anterior mantle (the pallial oviduct), and
are born free-living. Some viviparids in the genus
Campeloma, especially more northern populations, are
parthenogenetic (Van Cleave and Altringer, 1937; Vail,
1978; Johnson, 1992).
Pulmonates, in contrast, are all monoecious. The
basic components of the pulmonate reproductive system are shown in Figure 4. Sperm and eggs are produced in the ovotestis and exit via a common hermaphroditic duct in all pulmonates except ancylids, which
have two openings and are thus obligate cross-fertilizers. The albumen gland adds protein and nutrients to
the egg. Eggs are either fertilized in the hermaphroditic
duct by the same individual’s sperm, or by sperm from
another individual near the junction of the hermaphroditic duct and the oviduct. The external egg membranes
are then secreted in the oviduct. Eggs are laid in gelatinous egg cases and attached to plants, submersed
wood, other snails or rocks. Males have an introvertible penis, and fertilization is internal. Duncan (1975)
gives a more detailed discussion of egg and sperm formation, copulation and fertilization, and egg capsule
deposition.
Although pulmonates are simultaneous hermaphrodites, most species out-cross when possible. Pulmonates that “self” usually mature at later ages and
have lower fecundity. For example, Physella heterostropha suffers a reduction of 65% in fecundity
when not allowed to out-cross (Wethington and Dillon,
1993). Snails act as males or females based on the
amount of stored sperm that they possess (Wethington
and Dillon, 1996). Similarly, the African snail Bulinus
globosus suffers a 50% reduction in fecundity, and an
18% reduction in hatching rate, under obligate selffertilization, indicating a significant genetic load of recessive lethal genes (Jarne et al., 1991). In other
physids (Eileen Jokinen, personal communication), the
male system is larger in juveniles than in adults, and
protandry (a form of sequential hermaphroditism
where individuals switch sex from male to female as
they age and increase in size) may occur.
With the costs to fitness indicated in the preceding
paragraph, one might wonder why hermaphroditism
occurs in pulmonates. Most pulmonates are slow-moving and go through seasonal “bottlenecks” (precipitous
declines in density). The chances of finding a mate in
such situations are small, providing a selective advantage for monoecy. Pulmonates are dispersed passively
as spat trapped in mud on birds feet (Boag, 1986 and
references therein), and solitary, monoecious immigrants have an obvious advantage. The cost of inbreeding depression is evidently less than not being able to
reproduce at all. Parthenogenesis (being able to reproduce without males) may have a similar adaptive value
for prosobranch snails that are isolated in small headwater streams (Vail, 1978).
2. Digestive System
Food is brought into the mouth by rasping movements of the radula, a filelike structure (Fig. 5) resting
on a cartilage (the odontophore) to which muscles
which extend and retract the radula are attached.
When the radula is extended, it contacts the substratum, and algal particles are scraped off when retractors
pull the radula back into the mouth. The radula may
also pulverize food particles by grinding them against
the roof of the mouth. A long esophagus leads to the
10. Mollusca: Gastropoda
301
FIGURE 5
Scanning electron microscope photographs of radular teeth of (A) the lymnaeid, Pseudosuccinea
columella and (B) the physid, Physella vernalis (from Kesler et al., 1986). Note the lymnaeid has fairly large
teeth with few cusps. These teeth, along with large, cropping jaws in the buccal mass, a grinding gizzard
equipped with sand, and high cellulase levels allow it to specialize on filamentous green algae (Kesler et al.,
1986). Physella has many, small teeth with long cusps, arranged in chevron-shaped rows, useful for piercing
detritus and attached bacteria.
stomach, located in the visceral mass. Some gastropods
possess a specialized crop where sand grains further
abrade food particles. Digestive enzymes are produced
by the digestive gland, the hepatopancreas. Considerable digestion also occurs intracellularly in the hepatopancreas. Snails are one of the few animal groups
to possess and possibly synthesize cellulases (Kesler,
1983; Kesler et al., 1986), that degrade the algal cell
walls.
3. Respiratory and Circulatory Systems
The respiratory system differs radically between
prosobranchs and pulmonates. Freshwater prosobranchs
have a single ctenidium or gill (Aldridge, 1983). The
ctenidium, usually in the mantle cavity, has leaflike triangular plates richly supplied with blood vessels. Oxygen
poor blood passes across the plates in the opposite direction to oxygen-rich water currents (generated by ctenidal
cilia). This countercurrent mechanism assures diffusion
of oxygen from water into the blood. Pulmonates lost
their gill during their intermediate terrestrial phase, and
have a vascularized pocket in the mantle used as a lung
(hence their name). The opening of the lung is the pneumostome. Pulmonates either rely on surface breathing or
have a limited capacity for oxygen transfer across their
epithelial tissues (McMahon, 1983). Some physids and
lymnaeids fill the mantle pocket with water and use it as
a derived gill (Russell-Hunter, 1978). Ancylids and
planorbids have re-adapted further to aquatic conditions
by using a conical extension of epithelium as a gill, and
planorbids also have a respiratory pigment, hemoglobin,
which increases the efficiency of oxygen transport
(McMahon, 1983).
4. The Excretory System
Gastropods have a permeable epidermis and are
subject to osmotic inflow of water from their hypo-osmotic surroundings. Thus, they must pump out excess
water in their urine. The gastropod coelom is little
more than a small cavity (pericardium) surrounding the
heart. The coelomic fluid is largely a filtrate of the
blood, containing waste molecules, such as ammonia,
that are filtered across the wall of the heart. Additional
wastes are actively secreted into the coelom by the
walls of the pericardium. The coelomic fluid then enters a metanephridial tubule (the coelomoduct) where
selective resorption of salts and further secretion of
wastes occurs. The urine is then discharged into the
mantle cavity. Freshwater gastropods excrete nitrogen
both as ammonia and as urea. Ammonia is adaptive in
aquatic environments because, although it is toxic, it is
extremely soluble and readily diffuses away. Pulmonates often produce urea which is better in terrestrial situations, or during hibernation or estivation, because it is relatively nontoxic and can be stored in the
blood until able to be excreted (McMahon, 1983). Further discussion of excretion can be found in Martin
(1983) for gastropods in general and in Machin (1975)
for pulmonates.
C. Environmental Physiology
Pulmonates usually face wider extremes of temperature variation than do most prosobranchs (McMahon, 1983). Most temperate pulmonates, for example,
can withstand temperatures near 0°C for several
months, whereas tropical pulmonates can withstand
302
Kenneth M. Brown
temperatures near 40°C for extended periods. This is
undoubtedly adaptive because of the seasonal and diurnal variation in temperature in many pond habitats.
Pulmonates better regulate changes in metabolic rate
with changing temperatures than do prosobranchs. For
example, Q10 values for 18 pulmonate species were less
(averaging 2.2) than in 13 prosobranch species (averaging 2.8) (McMahon, 1983). As the average temperature
increases, snails grow faster and reproduce at an earlier
age, with more generations per year. For example, the
subtropical pulmonate Physella cubensis grows slowly
and will not mature at 10°C (Thomas and McClintock,
1990), but hatching times and ages at maturity decrease, and growth rates increase as laboratory temperatures are increased to 30°C.
Increasing water temperature is also the cue for onset of reproduction in many temperate pulmonates.
The ability of pulmonates to reproduce in cold water
allows adults to breed early in the spring and juveniles
to grow rapidly to adult size before the end of the summer. Pulmonates can secrete a mucus covering over the
aperture, called an epiphragm, to retard moisture loss
during dry periods (Boss, 1974; Jokinen, 1978).
In terms of adaptation to hypoxia, pulmonates: (1)
tolerate greater variation in dissolved oxygen than do
prosobranchs (possibly because prosobranchs rarely
experience hypoxia in the littoral zones of lakes or the
fast flowing rivers they are common in); and (2) also
regulate oxygen consumption at varying levels of dissolved oxygen better than prosobranchs, which are
“oxy-conformers” (Brown et al., 1998). Pulmonates
apparently withstand lower oxygen tensions by either
TABLE I
surface breathing or reliance on anaerobic metabolism
(McMahon, 1983).
Approximately 45% of freshwater gastropods are
restricted to waters with calcium concentrations greater
than 25 mg / L, and 95% to levels greater than 3 mg / L.
Although it may not take much energy to absorb calcium from water, assuming adequate water hardness, it
may be energetically costly to secrete calcium into the
shell against an electrochemical gradient. For example,
in the northeastern United States (Jokinen, 1987),
Aplexa elongata, Helisoma (Planorbella) trivolvis, and
all lymnaeids except Fossaria are limited to calciumrich waters. McMahon (1983) and Lodge et al. (1987)
discuss the degree of calcium regulation, and the relationship of external calcium level to shell thickness and
growth. Shell accretion may lag behind rapid tissue
growth in eutrophic habitats, producing thinner shelled
individuals (McMahon, 1983).
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
Although widespread in Asia, pleurocerids reach
their greatest abundance in the rivers and streams of
the southeastern United States (Table I). Most are fairly
large snails that are quite common in rocky riffles or
shoals in both headwaters and large rivers. However,
many species have been lost to stream alteration by impoundments, and present-day diversity has decreased
dramatically (see Section III.E). Females possess an egg
laying sinus on the right side of the foot (Dazo, 1965).
The Diversity of Freshwater Snail Families in North Americaa
Subclass
Family
Number of
genera
Prosobranchia
Ampullaridae
Bithyniidaec
Hydrobiidae
Micromelaniidae
Neritinidae
Pleuroceridae
Pomatiopsidae
Thiaridaec
Valvatidae
Viviparidae
Acroloxidae
Ancylidae
Lymnaeidae
Physidae
Planorbidae
2
1
28
1
1
7
1
2
1
5
1
4
9
4
11
Pulmonata
a
Data compiled from Burch (1989) and Neves et al. (1998).
N, North; E, East; W, West; S, South and U, ubiquitous.
c
Introduced.
b
Number of
species
4
1
152
1
1
156
6
3
11
29
1
13
58
43
47
Area of
greatest diversityb
SE
NE
U
SE
SE
SE
SE, W
S
U
E
W, NE
U
N
U
U
10. Mollusca: Gastropoda
Shell anatomy is used to classify genera, although many
species show considerable variation in shell characters.
The shells of pleurocerids are solid and the aperture
may bear a canal anteriorally. The operculum is corneous and paucispiral.
Viviparids are worldwide in distribution, and these
large snails are fairly diverse (Table I) throughout the
eastern states and Canadian provinces. Campeloma,
Lioplax, and Tulotoma are endemic to North America.
Although the first two genera are widespread, Tulotoma was once considered extinct and has only recently
been rediscovered in the Coosa River in Alabama,
where this filter feeding, brooding snail is common under rocks in shallow water shoals in large tributaries
(Hershler et al., 1990). Viviparus is quite common in
rivers and lakes throughout eastern North America.
Another large viviparid Cipangopaludina chinensis was
introduced to the United States in the 1890’s and has
now spread throughout the United States, especially in
hard-water lakes with sandy or muddy substrates (Jokinen, 1982).
Ampullarids are a tropical, mostly amphibious
family with a mantle cavity provided both with a gill
and a lung. The two genera present in Florida, Pomacea and Marisa, are quite large snails (50 – 60 mm).
Marisa has also been introduced to rivers in central
Texas.
The Neritinidae are a marine, tropical group. A
few species have invaded estuarine and freshwater
habitats, for example Neritina reclivata in Florida,
Georgia, Alabama, and Mississippi. The paucispiral,
calcareous operculum has a pair of projections which
lock the operculum against the teeth on the aperture,
providing a stronger defense.
Of the 20 species of valvatids in the northern hemisphere, 11 are found in North America (Table I),
mostly in lakes in northern states. Valvatids are egglaying hermaphrodites, with a single, featherlike gill
carried on the left side, and a pallial tentacle carried on
the right side of the shell as the animal crawls. Valvatids have small (approximately 5 mm diameter) dextral shells, with a corneous, slightly concave and thin,
multispiral operculum. The spire is only slightly elevated, and the shells are sometimes carinated.
Hydrobiids are extremely diverse and exist in
freshwater, brackish, and marine habitats. They are
small but have a thick shell which protects them from
fish predation, and they dominate the gastropod assemblages of many northern lakes (Brown, 1997). There
are 103 genera worldwide (Burch, 1989), and they are
diverse in North America (Table I). In fact, hydrobiids
may be the most diverse group of freshwater snails, as
58 new species in the genus Pyrgulopsis were recently
described from small springs in the southwestern
303
United States (Hershler, 1998). Although these populations often reach high densities, many species are endemic to only a few springs, increasing chances of extinction. The genus Fluminicola, which is common in
rivers in the northwestern United States, is also quite
diverse (Hershler and Frest, 1996). The hydrobiids
have small, dextral shells with a paucispiral operculum.
Because of the similarity of their shells, the structure of
the verge (penis) is used in classification.
The six North American pomatiopsids are similar
in general anatomy to the hydrobiids. Pomatiopsids
are, however, amphibious and are often found inhabiting stream banks several meters above the water’s surface, while hydrobiids are truly aquatic (Burch, 1989).
Pomatiopsid systematics are discussed by Davis (1979).
Thiarid females are parthenogenetic, brooding eggs
in a pouch in the neck region, which opens on the right
side. The similar-shelled pleurocerids, discussed above,
are on the contrary dioecious and oviparous.
Ancylids have a worldwide distribution, and all
possess a simple cone-shaped shell. In North America,
they have reached moderate diversity (Table I). Ancylids have sinistral shells, with the apex inclined
slightly to the right, and the gill (pseudobranch) and
many of the internal organs opening on the left side of
the body. Their streamlined shape allows them to colonize fast-flowing streams, where they are common on
rocks or macrophytes, although some species are also
common in lentic environments.
The family Acroloxidae occurs mainly in Eurasian
lakes and ponds. Only one species of Acroloxus occurs
in the United States, in Colorado and southeastern
Canada. Since the apex in Acroloxus is tipped to the
left, the aperture is considered to be dextral, unlike the
ancylid limpets.
Lymnaeids are worldwide in distribution, and are
the most diverse pulmonate group in the northern
United States and Canada (Table I). Lymnaeids have
broad triangular tentacles, and lay long, sausageshaped egg masses. Lymnaeids have fairly uncomplicated, large teeth on their radula (Fig. 5), which are
useful for cropping long strands of filamentous
algae(Kesler et al., 1986). One group of lymnaeids,
found along the Pacific coast of North America, has
limpet-shaped shells, but they are larger than ancylids.
Physids also have a worldwide distribution and are
ubiquitous in North America (Table I); few aquatic environments lack physids, because of their ease of introduction. Their shells are small, sinistral, with raised
spires. Their tentacles and foot are slender and they
have fingerlike mantle extensions (except for the genus
Aplexa) and lay soft, crescent-shaped egg masses. Their
radular teeth are smaller and more complicated in
shape than in the lymnaeids (Fig. 5), and they are
304
Kenneth M. Brown
better at harvesting more tightly attached periphyton
species like diatoms, or in feeding on detritus (Kesler
et al., 1986). Their rate of crawling is much more rapid
than most gastropods, and this along with their early
age at maturity and high fecundity explains why they
are so wide-spread (Brown et al., 1998).
Planorbids are widespread and fairly diverse snails
(Table I), and range in size from minute (1 mm) to
quite large (30 mm) in North America. They possess
hemoglobin as a respiratory pigment, sometimes giving
the tissue a red hue. Their egg cases are flat and circular, with harder membranes than those of the lymnaeids or physids. Planorbids are considered closely related to the ancylids (McMahon, 1983).
B. Reproduction and Life History
Freshwater snails are extremely interesting because
of the variety of observed life-history patterns. For
example, freshwater pulmonates are oviparous (egglaying) hermaphrodites, and are usually annual and
semelparous (i.e., they reproduce once and die). On the
other hand, almost all prosobranchs are dioecious and
are often iteroparous, with perennial life cycles. Prosobranchs can be oviparous or ovoviviparous (RussellHunter, 1978; Calow, 1978, 1983; Brown, 1983). Annual species have essentially a 1 year life cycle, whereas
perennials often live and reproduce for 5 years or more.
Russell-Hunter (1978) and Calow (1978) have classified life histories for freshwater gastropods (Fig. 6). At
one end are annual adults that reproduce in the spring
FIGURE 6
The variety of life cycles in freshwater snails. Curves
represent the growth of individual cohorts (in millimeters of shell
length), circles represent size at maturity while triangles represent appearance of egg cases in samples. Panel A represents annual semelparity, and B – F represent increasing numbers of cohorts per year,
while panel G represents perennial iteroparity (from Calow, 1978).
SP, spring; S, summer; F, fall and W, winter.
and die (e.g., there is complete replacement of generations). Most pulmonates belong to this group, including species from the genera Lymnaea, Physella, and
Aplexa (the original studies are listed in Calow, 1978).
In the second category (Fig. 6B), reproduction occurs in
both spring and late summer with both cohorts surviving the winter, or (Fig. 6C) where there again is complete replacement of generations. In Figure 6D – F, there
are three reproductive intervals, with varying degrees of
replacement of generations. These would predominantly be populations in subtropical or tropical environments. Finally, there are populations that can truly
be considered perennial and iteroparous (Fig. 6G); most
are prosobranchs. For example, the pleurocerid genus
Elimia has life cycles lasting from 6 to 11 years in Alabama steams, with several cohorts overlapping at any
one time (Richardson et al., 1988; Huryn et al., 1994).
Adults reproduce in the spring and summer and are
more common in areas of low current velocity, while juveniles are common in the fall in high flow areas
(Huryn et al., 1994; Johnson and Brown, 1997).
Jokinen (1985) also found Russell-Hunter and
Calow’s life-cycle categories useful in describing lifecycle patterns in twelve species of gastropods inhabiting a
small Connecticut lake. Four species, Amnicola limosa,
Helisoma anceps, Helisoma (Planorbella) companulatum and Laevapex fuscus, had simple, annual patterns.
Four species had at least two breeding seasons a year,
with varying replacement of cohorts, including Pseudosuccinea collumella, Planorbula armigera, Promenetus
exacuous, and Gyraulus deflectus. Three species, Fossaria modicella, Gyraulus circumstriatus, and Physella
ancillaria, had continuous breeding throughout the
field season. The prosobranch Campeloma decisum
was the only species with a perennial reproductive pattern. Jokinen (1985) suggested that divergence in life
history patterns between taxonomically similar species
lessened competition, by allowing some species to specialize on detritus (common in the spring), and others
periphyton (common in the summer).
Marine snails have enormous fecundity, but individual eggs are extremely small. Most marine snails are
prosobranchs, and freshwater prosobranchs (their descendants) probably have relatively small eggs as a result. In fact, viviparids and thiarids may have evolved
ovoviviparity (and in some cases the production of
larger embryos) to cope with the much more unpredictable conditions in freshwaters (Calow, 1978,
1983). Similarly, the loss of the planktonic veliger, and
shortening of the developmental period in oviparous
freshwater snails has been attributed to the more variable physicochemical conditions (Calow, 1978).
Reproductive effort (percent of energy devoted to
reproduction) is lower in iteroparous freshwater snails
10. Mollusca: Gastropoda
than in semelparous ones (Browne and Russell-Hunter,
1978). For example, pulmonate families are semelparous with relatively high reproductive output (Brown,
1983). Pulmonates also reproduce at smaller sizes and
earlier ages, produce more eggs, have larger clutch sizes,
greater shell growth rates, shorter life cycles and smaller
final shell sizes than viviparid prosobranchs.
The reduced fecundity but increased parental care
found in viviparids (vs semelparous pulmonates) increases offspring survival. Life tables for Viviparus
georgianus (Jokinen et al., 1982) and for the pulmonate Lymnaea elodes (Brown et al., 1988) indicate
that survival to maturity is indeed much less than 1%
in all of the pulmonate populations, but over 40% in
the ovoviviparous prosobranch. Prosobranchs are also
sexually dimorphic in life-history patterns. Female viviparids live longer (males usually survive for only one
reproductive season) and reach larger sizes (Van Cleave
and Lederer, 1932; Browne, 1978; Jokinen, 1982; Jokinen et al., 1982. Pace and Szuch, 1985; Brown et al.,
1989).
Numerous studies have implicated periphyton productivity (Eisenberg, 1966, 1970; Burky, 1971; Hunter,
1975; Browne, 1978; Aldridge, 1982; Brown, 1985) in
determining voltinism patterns, growth rates, fecundity,
and gastropod secondary production (see review in Russell-Hunter, 1983). Other important factors include water hardness and temperature. For example, populations
of Elimia in limestone-substrate streams in Alabama
have greater annual production than populations in relatively impermeable slate or sandstone-substrate streams
(Huyrn et al., 1995), both because of the higher alkalinity and greater buffering of low temperatures in the winter by ground water. Similarly, populations of Lymnaea
stagnalis in Canada often take several seasons to complete their life cycle (Boag and Pearlstone, 1979) while
populations are annual in the warmer waters of Iowa
(Brown, 1979).
Lymnaea peregra in wave-swept habitats in English
lakes have r-selected life-history traits (e.g., early reproduction and high reproductive output) in comparison
to populations in less-harsh habitats (Calow, 1981).
However, other studies of life history variation in molluscs do not agree as well with the predictions of r- and
K-theory (see discussion in Burky, 1983).Transplant
studies, where individuals from separate populations
are reared in a common environment, have usually indicated that environmental effects on life histories are
much more important than genetic differences between
populations. For example, populations of Lymnaea
elodes reared in more productive ponds lay nine times
as many eggs, have an annual versus a biennial reproductive cycle, and reach larger individual sizes (Brown,
1985).
305
Although genetic polymorphism has been studied
using gel electrophoresis in terrestrial pulmonates and
freshwater prosobranchs more than in aquatic pulmonates, freshwater snails still appear to have levels of
genetic polymorphism intermediate to terrestrial and
marine species (Brown and Richardson, 1988). Terrestrial snails inhabit patchily-distributed microclimates
which increase chances for low population densities
and self fertilization, resulting in little genetic polymorphism within populations. Freshwater snails decline to
low densities because of seasonal bottlenecks and thus
may also self. Marine environments are less seasonal,
and many marine snails have planktonic larvae, facilitating gene flow and increasing polymorphism.
C. Ecological Interactions
1. Habitat and Food Selection, Effects on Producers
Slow moving, silty habitats are occupied by pulmonates or detritivorous prosobranchs such as viviparids, whereas fast current areas are dominated by
limpets or prosobranch grazers like pleurocerids (Harman, 1972). There is also evidence for habitat selection
on the species level. Campeloma decisum is positively
rheotactic (moves upstream) and aggregates at any barrier (e.g., logs, riffle zones, etc., Bovbjerg, 1952). Physella integra and Lymnaea emarginata prefer cobble
substrates with attached periphyton, while Helisoma
anceps and Campeloma rufrum prefer sand (Clampitt,
1973; Brown and Lodge, 1993). Most gastropods in
northern Wisconsin lakes, however, prefer periphyton
covered cobble over sand or macrophytes (Brown and
Lodge, 1993). Cobble, since it offers greater surface
area and is present year-round, develops a richer periphyton coating. Substrate selection may even occur on
a finer level: gastropods from an English pond prefer
periphyton from the macrophytes the snails were found
on in the field (Lodge, 1986). Snails often move among
habitats as well. Migrations occur to deeper water in
the fall in lakes and back to the littoral zone in the
spring (Cheatum, 1934; Clampitt, 1974; Boag, 1981).
Pond pulmonates on the other hand burrow into the
substrate with declining temperatures (Boerger, 1975).
Freshwater gastropods are herbivores or detritivores, but occasionally ingest carrion (Bovbjerg, 1968)
or small invertebrates associated with periphyton
(Cuker, 1983). Periphyton is easier to scrape, and contains higher concentrations of nitrogen and other limiting nutrients than macrophyte tissue (Russell-Hunter,
1978; Aldridge, 1983). For example, carbon to nitrogen (e.g., carbohydrate to protein) ratios are below
10.1:1, while macrophytes have ratios of 24.1:1
(McMahon et al., 1974). Algal and diatom remains
therefore dominate in the guts of snails (Calow, 1970;
306
TABLE II
Kenneth M. Brown
Feeding Preferences of Freshwater Snails
Feeding type
Family
References
Algivores
(scrapers)
Ancylidae
Lymnaeidae
Detritivores or
bacterial feeders
Neritinidae
Pleuroceridae
Viviparidae
Physidae
Planorbidae
Viviparidae
Bithyniidae
Viviparidae
Calow (1973a, b, 1975)
Bovbjerg (1968, 1975), Brown (1982), Calow (1970), Cuker (1983), Hunter (1980),
Kairesalo and Koskimies (1987), Kesler et al. (1986), Lodge, (1986)
Jacoby (1985)
Aldridge (1982, 1983), Dazo (1965), Goodrich, (1945)
Duch (1976), Jokinen et al., (1982)
Brown (1982), Kesler et al., (1986), Townsend (1975)
Calow (1973b, 1974a, b)
Chamberlain (1958), Pace and Szuch, (1985), Reavell (1980)
Tashiro (1982), Tashiro and Colman (1982), Brendelberger and Jurgens, (1993)
Brown et al. (1989)
Filter feeders
1973a, 1975; Calow and Calow, 1975; Reavell, 1980;
Kesler et al., 1986; Lodge, 1986; Madsen, 1992).
However, gastropods at high densities can exhaust periphyton and then consume macrophytes, suppressing
macrophyte species richness (Sheldon, 1987).
Feeding preferences for the freshwater gastropod
families are summarized in Table II. Lymnaeids are “microherbivores”, scraping algae and diatoms from rocks
or macrophytes, but grow more rapidly when animal
tissue is in their diet (Bovbjerg, 1968). For example, although Pseudosuccinea columella (a lymnaeid) is an omnivore, it still consumes more algae than the sympatric
Physa vernalis (Kesler et al., 1986). The lymnaeid also
possesses higher levels of cellulases, as well as a radula
and jaws well adapted for cropping algae (Fig. 5), and a
gizzard filled with sand that can macerate food. The
physid, a detritivore, lacks these adaptations.
Both the limpet family Ancylidae and the prosobranch family Pleuroceridae are also considered to
graze on periphyton (Table II). Ancylus fluviatilis selectively grazes diatoms, but the limpet has little effect on
periphyton communities, due either to adaptations of
algal and diatom species to grazing or to relatively low
limpet abundances (Calow, 1973a, b). Aldridge (1983)
concluded that pleurocerid grazers feed on periphyton
rather than macrophyte tissue again because of higher
levels of nitrogen.
The prosobranch Bithynia tentaculata both grazes
on periphyton and uses its ctenidium to capture phytoplankton that is consolidated into a mucous string
which loops from the mantle cavity to the mouth
(Brendelberger and Jurgens, 1993). Indeed, filter feeding may be more efficient than scraping as increasing
levels of phytoplankton shade out periphyton, explaining why this species has become so abundant in nutrient rich, eutrophic lakes in New York (Tashiro, 1982;
Tashiro and Colman, 1982).
Viviparus georgianus is a micro-algivore (Duch,
1976; Jokinen et al., 1982) or a detritivore (Pace and
Szuch, 1985). Most viviparids, however, are probably
detritivores or utilize bacteria associated with detritus
(Table II). As macrophytes decompose, nitrogen levels
increase, again increasing their value as food resources.
For example, Viviparus reaches extremely high densities
(from 151 to 608 individuals m2) in wooded streams
in Michigan (Pace and Szuch, 1985) and detritus-rich
bayous in Louisiana (up to 1,700 individuals m2, Brown
et al., 1989).
Both physids (Kesler et al., 1986) and planorbids
(Calow, 1973b, 1974a) prefer detritus (Table II). For
example, although widely-spread species like Physella
gyrina and Helisoma trivolvis did not prefer detritus
over periphyton in laboratory experiments, another
physid, Aplexa elongata, which is much more common
in wooded ponds with a rich detritus food base, preferred detritus (Brown, 1982).
Originally, snail algivores were considered indiscriminant grazers, taking all components of the periphyton (Hunter, 1980; Hunter and Russell-Hunter,
1983). However, limpets and planorbids are selective
(Calow, 1973a, b), and Lymnaea peregra grazes selectively on filamentous green algae (Lodge, 1985). Planorbis vortex ingests diatoms in greater quantities than
found in the periphyton, but is still predominantly a
detritivore.
Gastropods have the ability to locate macrophytes
through distant chemoreception (Croll, 1983). For example, Lymnaea peregra is positively attracted to Ceratophylum demersum because of dissolved organic materials excreted by the macrophyte (Bronmark, 1985b).
Similarly, Potamopyrgus jenkinsi orients toward both
plant and animal extracts (Haynes and Taylor, 1984),
while Biomphalaria glabrata either orients toward or
away from specific macrophytes (Bousefield, 1979).
Almost all experimental manipulations have indicated snail grazers can decrease periphyton standing
crops (Table III, see also review in Bronmark, 1989).
For example, Physella at high densities reduces algal
10. Mollusca: Gastropoda
307
TABLE III The Effects of Experimental Manipulations of Gastropods on Periphyton Biomass, Production and
Assemblage Structure
Decreased
algal
biomass?
Increased
algal
production?
Favored
adnate
species?
Group
Genus
Prosobranchs
Elimia or Juga
(N 11)
73%
17%
100%
Theodoxus (N 1)
Amnicola (N 1)
Physella (N 3)
Y
Y
100%
NO
N.M.
N.M.
Y
Y
100%
Promenetus (N 1)
Lymnaea,
Physella,
Helisoma
NO
Y
NO
Y
NO
Y
Pulmonates
Studies
Mulholland et al. (1983), Steinman et al. (1987),
Lamberti et al. (1987), Marks and Lowe (1989),
McCormick and Stevenson (1989), Hill and
Harvey (1990), Mulholland et al. (1991),
Tuchman and Stevenson (1991), Hill et al. (1992),
Rosemond et al. (1993)
Jacoby (1985)
Kesler (1981)
Doremus and Harman (1977), Lowe and Hunter
(1988), Swamikannu and Hoagland (1989)
Doremus and Harman (1977)
Hunter (1980)
Percentages refer to the number of studies noting an affect. For single studies: Y, effect; NO, no effect, N.M., not measured.
biomass by 97%, and richness by 66% (Lowe and
Hunter, 1988). In some cases, snails also increase algal
production, perhaps by decreasing total biomass and
lowering algal competition for light or nutrients, removing senescent cells, or increasing rates of nutrient
cycling. Pulmonate gastropods can also alter the quality (e.g., nitrogen-to-carbon ratios and chlorophyll a
levels) of periphyton (Hunter, 1980), as can prosobranchs in streams (Steinman et al., 1987). Although
low levels of grazing may stimulate production, higher
snail densities decrease both biomass and production
(McCormick and Stevenson, 1989; Swamikannu and
Hoagland, 1989). Grazing may not, however, have as
much of an impact on shaded streams, where light may
be the primary limiting factor (Hill and Harvey, 1990).
Even in unshaded steams there may be little overall effect of grazers, because the loss of the algal over-story
to grazing is compensated for by the competitive release and increased growth of adnate algae (Hill and
Harvey, 1990).
Snail grazers selectively remove larger filamentous
green algae, and leave smaller, adnate species behind
(Table III). Under slight gastropod grazing pressure, periphyton assemblages are dominated by filamentous
green algae, but more intensely grazed assemblages are
dominated by more tightly adhering or toxic species
such as cyanobacteria. Snail grazers may, in fact, indirectly facilitate macrophytes, if periphyton coverings
shade and limit macrophyte growth, and snails prefer
periphyton. For example, the growth of Ceratophylum
demersum increased when gastropod grazers were present (Bronmark, 1985b), although increased growth
also occurred when plants were exposed only to snail-
conditioned water, indicating increased nutrient recycling was a cause (Underwood, 1991). Similarly, when
sunfish depress snail abundances and increase periphyton abundance in fish enclosures, they may indirectly
depress macrophytes (Martin et al., 1992). The molluscivorous tench (a fish) can have similar cascading effects on European gastropods, periphyton, and macrophytes (Fig. 7). Thus, interactions between gastropod
predators, snails, periphyton, and macrophytes may be
very complex in natural systems.
2. Factors Regulating Population Size
One of the first demonstrations of population regulation under field conditions was with Lymnaea elodes
(Eisenberg, 1966, 1970). When the density of adult
snails in pens in a small pond was increased, adult fecundity declined, as did juvenile survival. With addition of a high quality resource, spinach, an increase in
the number of eggs per mass occurred. Evidently, the
availability of micronutrients in periphyton was the
crucial variable (Eisenberg, 1970). Brown (1985) provided additional evidence by transferring juvenile Lymnaea elodes among a series of ponds differing in periphyton productivity. There was an exponential increase
in juvenile growth rates with increasing pond productivity, and snails in the most productive pond laid nine
times as many eggs as snails in the two less productive
ponds. A number of field studies also provide indirect
evidence for the importance of resource abundance:
populations in more eutrophic habitats have more generations a year, more rapid shell growth, and lay more
eggs (Burky, 1971; Hunter, 1975; McMahon, 1975;
Eversole, 1978). Highly eutrophic sites may however
308
Kenneth M. Brown
FIGURE 7
Cascading (also called top-down) effects of a molluscivorous fish (the tench) on gastropods, periphyton and macrophytes,
based on a field manipulation (from Bronmark and Vermaat, 1998).
Note how snail numbers are depressed by fish predation (top),
and reduced gastropod grazing results in periphyton increases,
and macrophytes are shaded by increasing periphyton cover, and
decrease.
be detrimental to gastropods. For example, gastropod
diversity declined over a fifty year period as Lake
Oneida, New York, became increasingly eutrophic
(Harman and Forney, 1970). In lotic systems, manipulation of periphyton resources and the abundances of
pleurocerid grazers have indicated both that food resources can control snail density and size distributions,
and that the snails can, in turn, control their food resources (Hill et al., 1992; Rosemond et al., 1993).
Although less studied than the role of periphyton,
the parasitic larvae of trematode worms may also impact snail population dynamics and evolution (Holmes,
1983). The adult worm produces eggs that are expelled
in the feces of the final host (a vertebrate) and hatch
into an infectious larval stage called a miracidium. Depending on the parasite group, the miracidium either
penetrates the epithelium of the snail or is consumed as
an egg and hatches within the snail. Once inside the
snail host, miracidia asexually produce several stages
(redia and sporocysts) which eventually produce thousands of cercaria that infect the final host or another
intermediate host. Infections either accelerate or decelerate snail growth (Brown, 1978; Anderson and May,
1979; Holmes, 1983, see discussion in Minchella et al.,
1985). Immediately after infection or exposure, snail
egg production rates increase dramatically (Minchella
and Loverde, 1981), evidently reducing the eventual
costs to snail fitness. As parasites consume the ovotestis
and hepatopancreas, however, both growth and egg
production of “patent” snails (those infections in the final stage where cercaria are emerging from snails) drop
below that of uninfected snails (Minchella et al., 1985).
The role of trematodes in controlling snail populations is unclear, as prevalence (percentage of population infected) varies considerably (Brown, 1978;
Holmes, 1983). For example, prevalence in Lymnaea
elodes in Indiana ponds varied from as low as 4% to as
high as 49% (Brown et al., 1988). Prevalences were
higher in less productive ponds, evidently because food
limitation caused longer snail life cycles that increased
chances for snails to be located by miracidia. Life-table
models predicted that the number of offspring produced per adult in the next generation declined by 14
to 21% in parasitized populations of L. elodes.
Assemblages of trematode larval species within a
snail host can themselves be considered communities,
with relative abundances and types of biotic interactions dependent on the snail species, the type of final
host, and the microhabitat the snail occupies. For example, multispecies infections are common and interspecific competition rare in Physella gyrina, evidently
because it is quite mobile and reproduces continually
throughout the field season, traits which increase
chance of infection (Snyder and Esch, 1993). Helisoma
anceps is more sedentary and has a simple annual lifehistory pattern; its parasitic trematode community is
characterized by fewer multiple infections and a competitive dominance hierarchy among parasites. Regarding microhabitats, trematode larvae that infect snails
when eggs are consumed (and that have frogs as final
hosts) are more common in snails collected from shallow habitats in ponds (Sapp and Esch, 1994); in contrast, while larvae that infect snails as miracidia (and
10. Mollusca: Gastropoda
309
TABLE IV Comparison of Average Standing Stocks, Productivity, and Turn-over Times for Populations
of Pulmonate and Prosobranch Snails Reported in Russell-Hunter and Buckley (1983)
Subclass
Mean biomassa s.e.
Mean productionb s.e.
Mean
turnover timec s.e.
Pulmonata
Prosobranchia
0.98
4.64
5.71
3.56
98.0
385.3
0.50 (6)
1.80 (4)
2.44 (10)
0.51 (5)
9.46 (10)
33.5 (4)
(N) Number of species averaged.
a
g C / m2.
b
mg C / m2 per day.
c
days.
have waterfowl as final hosts) occur in all microhabitats, but only when the final hosts are common.
Trematode parasites and their snail hosts are also
extremely interesting from a co-evolutionary viewpoint
(Holmes, 1983; Minchella et al., 1985). Because invertebrates cannot easily acquire resistance to parasites,
frequency-dependent selection may operate to ensure
the fitness of any genotype less vulnerable to a particular trematode (Holmes, 1983). Parasites may also cause
a shift in investment of resources from costly reproduction to growth and maintenance and, thus, even increase survivorship of infected snails (see also Baudoin,
1975; Minchella et al., 1985). Because trematode larvae evolve surface antigens that mimic those of the
snail host, application of one evolutionary model,
called the “red queen” hypothesis, suggests that gastropods are constantly evolving new antigen phenotypes merely to stay ahead of their trematode parasites.
For example, the New Zealand prosobranch Potamopyrgus has both sexual morphs (able to produce genetically more variable offspring through recombination) and parthenogenetic morphs. The frequency of
sexual reproduction is positively correlated with trematode prevalence, as one would expect if sexual morphs
have an advantage (Lively, 1987). Sexual morphs are
also more common in shallow water, where waterfowl
(the final hosts) occur (Lively and Jokela, 1995). Infected snails are also more likely to forage during the
day than uninfected snails or gravid females, suggesting
parasites may modify snail behavior so they are more
likely to be consumed by waterfowl, to complete the
life cycle (Levri and Lively, 1996). Experimental studies
in New Zealand have suggested trematode populations
are locally adapted to better infect their own host populations than snails from different lakes (Lively, 1989).
Gene flow has not swamped such local adaptation,
even though rates of gene flow are greater among parasite populations than among their snail hosts (Dybdhal
and Lively, 1996), However, sexual reproduction may
not be favored in all snail – trematode systems. In
Campeloma, certain trematode metacercaria actually
feed on sperm, and parthenogenetic morphs thus have
an advantage (Johnson, 1992).
3. Production Ecology
Average standing crop biomass, productivity, and
turnover times (these terms are explained in the glossary) differ between prosobranchs and pulmonates
(Table IV, summarized from Russell-Hunter and Buckley, 1983). Standing crops are greater on the average
for prosobranchs than for pulmonates, although prosobranch populations studied to date have not included
genera with smaller individuals, such as Amnicola and
Valvata.
Even if pulmonates have lower standing stocks,
their rapid growth and short life cycles still result in
higher average production rates and shorter turnover
times (Table IV). For example, two pleurocerid grazers
in an Alabama stream, Elimia cahawbensis and E.
clara, have considerable biomasses of 2 – 5 g ash-free
dry mass (AFDM) per square meter, but slow growth
rates and long life cycles result in relatively low rates of
secondary production (0.5 – 1.5 g AFDM / m2). The
combination of high biomass and low production results in a low production to biomass ratio of 0.3
(Richardson et al., 1988). Prosobranch detritivores
may, however, have higher production rates. Viviparus
subpurpureus and Campeloma decisum, with high densities and short life cycles in Louisiana bayous, have
high standing crop biomasses (10 – 20 g AFDM / m2)
and production rates (20 – 40 g AFDM / m2 per year)
among the highest known for freshwater molluscs
(Richardson and Brown, 1989).
4. Ecological Determinants of Distribution
Water hardness and pH are often considered major
factors determining the distributions of freshwater
snails (Boycott, 1936; Macan, 1950; Russell-Hunter,
1978; Okland, 1983; Pip, 1986). For example, in the
acid-rain affected lakes of the Adirondack mountains
of New York, Jokinen (1991) found that snail diversity
declined with declining pH and increasing altitude.
310
Kenneth M. Brown
However, in lake districts with adequate calcium
(above about 5 mg / L CaCO3), or in the normal
(nonacidified) range of pH, relationships between
physico-chemical parameters and gastropod diversity
are less clear (Lodge et al., 1987; Jokinen, 1987). For
example, species in New York lakes overlap broadly in
the ranges of physico-chemical variables in which they
occur (Harman and Berg, 1971). Physico-chemical parameters, therefore, set the limits for gastropod distributions, but are not as important in explaining the relative abundance patterns and densities of gastropods in
most hard-water, circum-neutral lakes.
On a biogeographic scale, a factor determining
gastropod distributions is dispersal ability. Studies have
indicated diversity increases with the area of lakes and
ponds (Lassen, 1975; Browne, 1981; Bronmark,
1985a; Jokinen, 1987). Since immigration rates generally increase and extinction rates decrease with increasing habitat size, larger habitats, all else being equal,
usually support more species (MacArthur and Wilson,
1967). Jokinen (1987), following the ideas of Diamond
(1975), found that snail species were not distributed
randomly with increasing snail diversity. Five snails
were “high-S” species (e.g., those that occurred only at
the most diverse sites) including Valvata tricarinata,
Gyraulus deflectus, Laevapex fuscus, and two species
of Lyogyrus. Most gastropods were widely distributed
across sites (“C – D” tramps in Diamond’s terminology). The species with the greatest dispersal ability
(called “super tramps” by Diamond) were found only
at low diversity sites; these included the lymnaeid Pseudosuccinea collumella and the limpet Ferrissia fragilis.
After dispersal, successful colonization depends on
the presence of suitable substrates. For example, a significant relationship exists between the number of gastropod species and the number of substrates in lakes
and streams in New York, USA (Harman, 1972). In
fact, substrate preferences determined in the laboratory
are good predictors of the types of ponds where the
snails are common (e.g., algivores in open ponds, detritivores in wooded ponds; Brown, 1982). Gastropod diversity is also positively related to macrophyte biomass,
probably because macrophytes increase surface area for
periphyton colonization (Brown and Lodge, 1993).
Disturbance may also determine the assemblage of
snails. In temporary ponds, diversity is lowered by frequent drying; and in habitats that go hypoxic, diebacks
of macrophyte and gastropod populations will also occur (Lodge and Kelly, 1985; Lodge et al., 1987). Disturbance may also limit some species from areas such
as wave-swept shores, littoral zones of reservoirs, etc.
Similarly, in streams, current velocity may affect distribution and growth. Adult Elimia avoid high-flow areas, perhaps because increased flow makes movement
more difficult and lowers grazing rates or increases
metabolic rate, thus lowering adult size (Johnson and
Brown, 1997). Juveniles, because they are smaller and
can exploit the boundary layer, are actually more common in fast-flowing areas.
In large lakes, biotic interactions such as interspecific competition or predation, are important in determining gastropod diversity and abundance (Lodge
et al., 1987). In regards to competition, some evidence
suggests it is rare in freshwater snails. For example,
pulmonate pond snails in the midwestern USA overlap
little on food and habitat dimensions of the niche, although some species do compete (Brown, 1982). Other
studies also indicate differences in resource utilization.
For example, the European ancylid Ancylus fluviatilis
prefers diatoms and is found on the top of cobble
where periphyton is abundant; whereas the co-occurring planorbid Planorbis contortus is a detritivore and
occurs more often under stones, where detritus accumulates (Calow, 1973a, b, 1974a, b). Similarly, Pseudosuccinea columella and Physa vernalis possess differences in their gut and radulae allowing coexistence in
New England ponds (Kesler et al., 1986). Furthermore,
niche partitioning may also be facilitated by differences
among snails in enzymatic activity (Calow and Calow,
1975; Brendelberger, 1997) or radular morphology
(Barnese et al., 1990).
However, there is some indirect evidence for competition. First, there are often fewer coexisting congeners in field samples than would be expected, based
on mathematical simulations (Dillon, 1981, 1987). Second, competition has been inferred from changes in relative abundance of gastropod species through time,
that is, by apparent competitive exclusion (Harman,
1968). Third, niche overlap can, in fact, be higher in
more diverse snail assemblages found in large lakes.
For example, the abundances of gastropod species are
often positively associated with high-diversity macrophyte beds in lakes, and experiments show most species
have similar preferences for macrophytes (Brown,
1997). Finally, pulmonates are common in ponds or in
vegetated areas of lakes, while prosobranchs are rare in
ponds and common in lakes and rivers. One explanation could be competitive exclusion of pulmonates
from lakes by prosobranchs, or exclusion of prosobranchs from ponds by the same mechanism. However,
additional explanations include greater vulnerability of
prosobranchs to hypoxic conditions in ponds, poorer
dispersal abilities of prosobranchs, or the fact that thinshelled pulmonates are more vulnerable to shell-crushing fish common in lakes or rivers (Brown et al., 1998).
Indeed, Lodge et al. (1987) argue that predators determine the composition of gastropod assemblages in
lakes. Indirect evidence includes the fact that gastropods
10. Mollusca: Gastropoda
have a number of anti-predator adaptations. These include thick shells to protect against shell-crushing
predators (Vermeij and Covich, 1978; Stein et al., 1984,
Brown and DeVries, 1985; Saffran and Barton, 1993;
Brown, 1998), as well as escape behaviors such as shaking the shell or crawling above the water to protect
against shell-invading invertebrate predators (Townsend
and McCarthy, 1980; Bronmark and Malmqvist, 1986;
Brown and Strouse, 1988; Alexander and Covich, 1991
a, b; Covich et al., 1994). Molluscivores are common in
lakes and rivers. For example, the pumpkinseed sunfish,
Lepomis gibbosus, and the redear or shell-cracker sunfish, Lepomis microlophus, specialize on gastropod prey
and have pharyngeal teeth adapted to crush shells.
Crayfish will select snails instead of grazing on macrophytes if given a choice (Covich, 1977), using their
mandibles to chip shells back from the aperture and selecting species with thinner shells (Saffran and Barton,
1993; Brown, 1998).
Although Osenberg (1989) argued that lentic snail
assemblages are limited by food resources, most experimental evidence supports a strong role for predators in
determining snail diversity and abundance (Table V).
For example, the central mud minnow can significantly
lower the density of relatively thin-shelled snails in permanent ponds. When fish densities were manipulated
in pens, the number of eggs and juveniles of Lymnaea
TABLE V
311
elodes was significantly less in the presence of fish. The
small, gape-limited fish feed on eggs and juveniles and
may restrict Lymnaea elodes from permanent habitats
such as large marshes or lakes. Pumpkinseed sunfish
also strongly prefer large, weak-shelled gastropod
species in laboratory experiments, and thin-shelled
species also decline dramatically in pumpkinseed enclosures in lakes (see references in Table V). Thus, most
thin-shelled pulmonates should occur in lakes only in
macrophyte beds, where they have a refuge from visual
predators; and sandy areas should be dominated by
thicker shelled pulmonates like Helisoma or by prosobranchs. Indeed, gastropod distributions among habitats within Indiana and Wisconsin lakes do follow these
patterns (Lodge et al., 1987; Brown and Lodge, 1993;
Lodge et al., 1998). In fact, fish predators may indirectly benefit poorer gastropod competitors by preferentially removing dominant species. Lake trout, for example, preferentially consume larger lymnaeids, releasing
valvatids from competition in arctic lakes (Hershey,
1990; Merrick et al., 1991). However, experimental manipulations suggest such indirect effects do not extend
through four trophic levels. Piscivores (top predators
feeding on fish) in Swedish lakes cannot depress predators of snails to the extent that interactions cascade
down the food web to facilitate snails which negatively
impact periphyton (Bronmark and Weisner, 1996).
Summary of Experimental Field Manipulations Testing the Role of Snail Predators in Controlling Their Prey
Study
Prey
Predator
Conclusions
Brown and
Devries 1985
Sheldon 1987
Lymnaea
elodes
assemblage
mud minnows
decreased abundance of eggs and juveniles of thin-shelled L. elodes
sunfish
Kesler and
Munns 1989
Osenberg 1989
assemblage
belastomatid bugs
removal of fish increased snails which decreased macrophytes
through herbivory
decreased snail abundances in New England pond
assemblage
littoral sunfish
Hershey (1990),
Merrick et al.
(1991)
Martin et al.
1992
Bronmark
et al. 1992
Bronmark 1994
Lodge et al. 1994
assemblage
lake trout
assemblage
assemblage
assemblage
L. microlophus
L. macrochiras
pumpkinseed
(L. gibbosus)
Tench
O. rusticus
Daldorph and
Thomas 1995
Bronmark and
Weisner 1996
assemblage
sticklebacks
fish decreased snails, increased periphyton, decreased macrophytes
crayfish decreased snails and macrophytes, had no effect
on periphyton
fish decreased thin-shelled snail spp., increased periphyton
assemblage
piscivores &
molluscivores
molluscivores did decrease snails, but piscivores did not control
molluscivores and increase snails indirectly
assemblage
Assemblage refers to a natural community of gastropods.
fish controlled only rare, large snail spp., competition between
snails considered more important
trout removed better competitor (L. elodes), which favored poorer
competitor, Valvata
fish decreased snails 10-fold, increased periphyton biomass 2-fold,
decreased algal cell size, decreased macrophytes.
fish decreased snails, increased periphyton, favored adnate algae
312
Kenneth M. Brown
Invertebrate, shell-invading predators may also
limit snail populations or cause shifts in gastropod relative abundances. Although some leeches, such as Nephelopsis obscura, have fairly low feeding rates (Brown
and Strouse, 1988), crayfish can consume up to or over
a hundred snails per night. The crayfish Orconectes
rusticus significantly reduced snail abundances in enclosure experiments in Wisconsin lakes (Table V), and
an interlake survey indicated that snail abundances
were negatively correlated with crayfish catches. Crayfish also affect gastropod habitat selection: even though
cobble habitat is preferred by many snails (see earlier
discussion), it also provides refugia from fish predators
for crayfish. Crayfish predation overrides the effects of
rich food resources (Weber and Lodge, 1990).
Crayfish are also quite selective molluscivores.
They prefer thinner-shelled gastropods (Saffran and
Barton, 1993; Brown, 1998), and often prefer juvenile
snails which have shorter handling times and higher
profitability unless these more vulnerable prey are protected by macrophytes (Nystrom and Perez, 1998).
Crayfish predators also shift size distributions of Physella virgata upwards in Oklahoma streams. This may
be due both to size-selective predation and to the diversion of energy of the snail from reproduction to growth
in order to reach a size-based refuge from predation
(Crowl, 1990; Crowl and Covich, 1990). Interestingly,
only snails in vulnerable size classes respond by crawling out of the water after predation by crayfish has occurred on conspecifics (Alexander and Covich, 1991b).
Other invertebrate predators may also be important.
For example, belostomatid bugs can eat as many as five
snails per bug per day in the laboratory (Kesler and
Munns, 1989).
Fish predators may alter snail foraging behavior as
well. In experiments, Physella selects refugia in the
presence of sunfish predators, only leaving when sufficiently starved (Turner, 1997). This behavioral interaction may provide an additional explanation for why
periphyton biomass increases when fish predators are
present. For example, use of open habitats by snails decreases with increasing predation risk from sunfish
(Fig. 8), with the result that periphyton biomass also
increases (Turner, 1997). Interestingly, such indirect facilitation of periphyton does not occur upon experimental addition of crayfish to enclosures, probably because crayfish are also omnivorous and graze on
periphyton (Lodge et al., 1994).
5. Suggestions for Further Work
Studies of the production ecology of smaller prosobranchs, as well as snail populations in other areas besides the northeastern United States are still needed.
Questions still exist on subjects such as:
FIGURE 8
Indirect effect of mortality from fish predation on snail
grazing rates. As more crushed snails were added to experimental
pools (predation risk perceived by snails, x-axis), snails used refuges
more, resulting in reduced use of open areas (left y-axis). Note how
periphyton biomass (ashfree dry mass or AFDM, right y-axis)
increased in open area as a result of reduced snail grazing (after
Turner, 1997).
1. How pulmonates are physiologically adapted
to relatively ephemeral or hypoxic aquatic
habitats, including studies of differences in
metabolic pathways and nitrogen excretion
(McMahon, 1983).
2. The relative roles of the physiological and ecological constraints to shell versus tissue growth,
given the remarkable intraspecific variation in
shell structure and composition in freshwater
gastropods.
3. Physiological bases of feeding preferences (such
as relative cellulase activities, etc.).
4. Micro-preferences (e.g., preferences for certain
algae or diatom species), for example, whether
detritivores are consuming leaves or more nutritious bacteria or fungi.
5. The nutritional quality or toxicity of different
food types and their role in preferences by
gastropods.
Further research on abiotic and biotic factors influencing snail assemblages is also necessary. Abiotic factors such as pond drying, anoxia, floods, or macrophyte die-offs could explain much about gastropod
population dynamics or species composition. Further
work is also necessary to determine whether the allopatric distributions of pulmonates and prosobranchs
are due to competition (see discussion in Brown et al.,
1998). Further work is needed as well on the relative
role of invertebrate predators like leeches, belostomatid
bugs, and scyomyzid fly larvae (Eckblad, 1973) in
structuring snail assemblages.
10. Mollusca: Gastropoda
313
D. Evolutionary Relationships
Marine prosobranchs are ancestral to freshwater
prosobranchs and terrestrial and freshwater pulmonates. Freshwater prosobranchs adapted to the dilute osmotic conditions of estuaries and then rivers,
and most modern families are widespread because their
adaptive radiations predated continental breakups and
drift. Davis (1979, 1982) and Clarke (1981) pointed
out that prosobranch dispersal is limited in most cases
to slow movement of adults along streams and rivers.
Such populations are more likely to become isolated,
promoting chances of speciation and adaptive radiation. Examples include the dramatic adaptive radiation
of hydrobiids (Davis, 1982) and pleurocerids in lotic
habitats (Burch, 1989; Lydeard et al., 1997).
In contrast, pulmonates, due to their greater rates
of passive dispersal as spat on birds and insects, have
more widespread distributions. For example, they are
widespread in the northern and northeastern states, as
well as in Canada (Harman and Berg, 1971; Clarke,
1981). Immigration rates to ponds average 0.8 and can
be as high as 9 species per year (Lassen, 1975; Davis,
1982). Pulmonates also predominate in shallow, more
ephemeral habitats less than 100 km2 in area and with
durations less than 103 years. Populations probably do
not exist long enough in such habitats for speciation to
occur, explaining why pulmonates are less speciose
than prosobranchs (Russell-Hunter, 1983; Clarke,
1981; Davis, 1982).
Pulmonates apparently evolved from intertidal
prosobranchs that relied less and less on aquatic respiration (Morton, 1955; McMahon, 1983). Modern estuarine pulmonates like Melampus may resemble these
ancestral species. The intermediate, terrestrial pulmonates lost the ctenidium, and gave rise both to modern terrestrial pulmonates (order Stylommatophora)
and the aquatic pulmonates (order Basommatophora).
Figure 9 illustrates a hypothetical phylogeny of aquatic
pulmonates based on progressive re-adaptation to
aquatic habitats (see discussion in McMahon, 1983).
Lymnaeids are in some cases still amphibious (e.g., occurring in temporary wetlands), while physids are intermediate in their adaptation to aquatic habitats
(some species come to the surface to breathe, while
others have filled the lung with water and can be found
in deeper lakes). Ancylids and planorbids have secondarily acquired more aquatic specializations such as the
evolution of secondary gills, and in the planorbids, hemoglobin.
Several phylogenetic trees for freshwater snails have
been published recently, using both classical morphological and recently developed molecular methods. Jung
(1992) used the more traditional characters of shell and
FIGURE 9 Hypothetical phylogeny for freshwater pulmonates
assuming increased adaptation through time to secondarily-invaded
aquatic habitats. The argument is that air-breathing lymnaeids are
limited to shallow water, while physids, which in some cases can fill
their lung with water and use it as a gill, are found in deeper water,
and ancylids and planorbids are the most specialized for an aquatic
life style, with secondarily derived epithelial gills and in the planorbids, hemoglobin.
soft anatomy to propose a cladogram (Fig. 10) for certain species of planorbids, using 65 characters. He
pointed out that systematic studies should not rely on
preserved specimens, since fixation alters soft anatomy.
In his phylogeny, Biomphalaria is considered more ancestral, and Planorbella trivolvis and Helisoma anceps
the most derived species.
Remigio and Blair (1997) published a phylogeny
for certain lymnaeid species, using populations from
both North America and Europe (Fig. 11). They considered molecular methods, such as sequencing ribosomal DNA in mitochondria, superior techniques in
comparison to other characters such as shell morphology, which often converge among genera. For example, based on molecular information, they considered
karyotypes with 18 chromosomes to be ancestral, and
16 chromosomes to be a derived character, unlike
FIGURE 10 Simplified cladogram for several planorbid species
studied by Jung (1992), based on shell and soft anatomy.
314
Kenneth M. Brown
FIGURE 12
FIGURE 11 Cladogram for a subgroup of the lymnaeid species
found in both North America and Europe, based on molecular data.
Simplified from Remigio and Blair (1997).
earlier studies. European and North American stagnicolids (S. elodes and S. palustris) were not considered
each other’s closest relatives, contrary to interpretations based on morphology. Stagnicola catascopium, S.
emarginata, and S. elodes, based on genetic divergence, could actually be conspecific taxa. On the other
hand, two populations of Lymnaea stagnalis (one in
Germany and the other in Italy) were genetically quite
divergent, indicating isolation by the geographic barrier of the Alps. The authors propose that lymnaeids
evolved in the late Jurassic to early Cretaceous on the
common continent Laurasia, and were separated into
distinct Palearctic and Nearctic groups at the start of
the Cenozoic (65 million years ago) when ancestral
species still had 18 chromosomes. The genus Stagnicola was probably extant at that time, explaining
why North American and European species are so
divergent.
Lydeard et al. (1997) used mitochondrial rDNA
sequences to study the evolutionary relationships of
representative prosobranch snails in the Mobile River
Basin in the southeastern United States (Fig. 12). This
basin may have once had the most diverse gastropod
fauna in the world, with 118 species and nine families.
Pleurocerids make up almost two thirds of the surviving fauna. Because of impoundments, the genus Gyrotoma, once found in shallow riffles in the Coosa River,
has gone extinct, along with 14 species of Elimia, and
11 species of Leptoxis. Lydeard’s phylogeny considers
14 prosobranch species (with Melanopsis as an out
group) and suggests Elimia and Pleurocera are monophyletic genera, but that Leptoxis is paraphyletic. The
remaining Elimia species in the Mobile Basin are genetically fairly similar, while the remaining three species of
Cladogram for a subgroup of species in several prosobranch genera in the Mobile River Basin, based on molecular data.
The greater divergence of leptoxid species suggests a polyphyletic
origin for this genus. Simplified from Lydeard et al. (1997).
Leptoxis are quite divergent in their mitochondrial 16S
rDNA sequence.
Further studies of genetic relationships among
pleurocerids found in the Mobile River Basin suggest
that earlier species descriptions, relying solely on shell
morphology, have overestimated the true diversity of
the system. For example, Lydeard et al. (1998), again
looking at 16S rDNA sequences, considered Elimia hydei basal to all other elimiads, and Elimia crenatella, E.
showalteri, E. fascians, and E. E. caelatura to be true
species. The low genetic divergence within another
clade led them to question the status of six speces:
Elimia carinocostata, E. gerhardtii, E. haysiana, E. alabamiensis, E. olivula, and E. cylindracea. Dillon and
Lydeard (1998) used allozyme polymorphism to study
relationships among Mobile Basin leptoxids. They argued that Leptoxis praerosa and L. picta were deserving of species status, but that L. taeniata and L. ampla
should be subsumed within L. picta. On the other
hand, Dillon and Ahlstedt (1997) used allozyme variation to show that Athernia anthonyi and Leptoxis
praerosa were different species, although adult shell
morphology is similar.
Several caveats about the evolution of freshwater
snails should be kept in mind (Davis, 1982). First, phylogenetic analyses, and therefore biogeographic studies,
are often limited by the poor systematic information
available on many groups, as well as the poor fossil
record (only shells are preserved, which often show
convergence). Second, a number of factors can explain
current distributions, as illustrated for hydrobiids and
pomatiopsids by Davis (1982): (1) phylogenetic events
such as centers of origin and adaptive radiation; (2) past
history events such as continental drift and geological
alteration of stream and river flow; (3) dispersal powers, and (4) ecological factors (see earlier discussion).
10. Mollusca: Gastropoda
E. Conservation Biology of Freshwater Snails
Freshwater snails, especially prosobranch species in
the southeastern United States, are facing rates of extinction similar to the more widely known case of
North American unionid bivalves (Lydeard and Mayden, 1995; Neves et al., 1998; Jenkinson and Todd,
1998). Freshwater snails account for 60% of all North
American molluscan species, and 61% of these snails
are in the Southeast. Historically, there were 118
species of snails in the Mobile River Basin, 96 species
in the Tennessee River, and 36 species each in the Cumberland and Apalachicola River basins. Thirty-eight of
the 118 species in the Mobile River Basin are known to
be extinct. The decline in diversity in the Mobile River
Basin varies from 33 to 84% in different drainages.
Nine species of gastropods are currently listed as
federally endangered (Bogan, 1999), although over 200
species from 11 families are candidates (69% of which
are in the Southeast). In the southeastern states alone,
two viviparids, 50 hydrobiids, 83 pleurocerids, nine
pulmonates, three ancylids and six planorbid species are
considered at risk (Neves et al. 1998). Many of the
same factors that have imperiled unionid bivalves are
responsible: impoundments which inundate shallow riffles and decrease oxygen and alter water temperature,
increased turbidity from agricultural activities, improper sewage treatment, and other pollution sources,
to name but a few. Organic enrichment, pesticides and
pathogens endanger about a quarter of the impaired
river-miles in North American rivers, while excess nutrients and siltation occur, respectively, in 37 and 45% of
the impacted river-miles (Neves et al., 1998). At the current time there is little governmental interest in managing imperiled snail populations, although the large,
knobbed-shell species, Io fluvialis, has been successfully
reintroduced to one river in its original range.
315
Lodge et al. (1994). In coarse sand, Ponar dredges are
the best alternative. In cobble, little recourse to direct
counts of given areas by visual search is available. One
can estimate densities by individually covering the rocks
with aluminum foil, and then estimating the area from
weight-to-area regressions for the foil.
The best sorting technique for gastropods is handsorting. Large adults can be removed visually, while
samples are washed through a graded series of sieves
(the smallest having a mesh of approximately 0.5 mm)
to remove mud but retain smaller snails. Samples are
then carefully backwashed from the sieves into shallow
water on flat white trays, the vegetation teased apart,
and the whole tray examined in a systematic fashion to
remove small gastropods and egg cases.
Temperate gastropods will grow well at 15 – 20°C,
while subtropical species grow better at 20 – 25°C. Any
hard substrate with a dense periphyton covering can be
added for food, although artificial foods such as lettuce, cereal, or spinach are sometimes used. Provide
food only as needed to avoid fouling containers. Detritivores should be fed leaf litter colonized with bacteria
and fungi (i.e., held for at least 2 weeks in a pond or
stream).
Avoid crowding of snails, as growth and reproduction are sensitive to density. An approximate rule is one
snail per liter. Water should be re-circulated through a
gravel or charcoal filter, or at least be changed weekly.
Conspecifics should be paired so that mating can occur.
Culturing of “weedy” species like physids is best done
at low temperatures to retard egg production, and constant removal of egg cases is necessary to prevent population “explosions”. Adequate lighting (with a 12L:12D
cycle) is necessary to promote periphyton growth in
aquaria (gro-lites®work well).
V. IDENTIFICATION OF THE FRESHWATER
GASTROPODS OF NORTH AMERICA
IV. COLLECTING AND CULTURING FRESHWATER
GASTROPODS
A number of sampling techniques exist (reviewed
in Russell-Hunter and Buckley, 1983), although some
are not quantitative. The least quantitative technique,
but one that often gives large numbers of individuals
and a good idea of species composition, is sweep netting with a net of 1-mm mesh (Brown, 1979, 1997).
In soft sediments or sand, quantitative samples can
be collected with an Ekman grab or a corer. When sampling macrophytes, the sampler must collect both plants
with attached snails and the substrate with any bottomdwelling species. Examples of such samplers are described in Gerking (1957), Savino and Stein (1982) and
The following key is modified from Burch’s (1989)
key for North American freshwater snails. Taxa are
keyed down to genera in all families except the diverse
hydrobiids. Readers interested in keying specimens
down to the species level should consult Burch (1989)
or keys addressing regional snail faunas (Harman and
Berg, 1971; Jokinen, 1983; Thompson, 1984; Jokinen,
1992).
After collecting specimens, first relax them with tricaine methane sulfonate, menthol crystals, polypropylene phenoxytol, or sodium nembutol and then preserve them in 70% ethyl alcohol. Preserved specimens
are better than dried shells, as possession of an operculum will be one of the determinations necessary.
316
Kenneth M. Brown
Collect a number of different-sized specimens, because
closely related genera sometime differ only in adult
shell size. Sizes reported in the key, following Burch
(1989), are small shells less than 10 mm, medium between 11 and 29, and large greater than 30 mm in
length. Size bars in keys are in millimeters. Refer to
the glossary for shell sculpture terminology like costae,
lirae, etc. In some cases, soft tissue characters are used
in the key (for example the structure of the verge, or
penis, located behind the male’s right tentacle, is important in hydrobiids). The shells of preserved individuals can be carefully crushed and removed, after living
snails are relaxed with any of the chemicals mentioned
above. Examine the verge under a dissecting microscope.
If examination of the radula is necessary, dissect
out the buccal mass (a muscular mass behind the
mouth that surrounds the radula) from a relaxed
specimen and place it in a 10% potassium hydroxide
solution to digest the tissue. After several hours, remove any remaining tissue and rinse the radula in
70% alcohol, stain it and mount it on a slide for examination under a compound microscope. However,
digestion with strong bases may damage the radula.
An alternative technique (Holznagel, 1998) is more
involved, but also provides a method to extract DNA
for phylogenetic work, and can be used with frozen or
alcohol-fixed specimens (but not those preserved in
formalin). First prepare stock solutions of NET buffer
and Proteinase-K. For the former, add 1 mL pH 8.0
Tris buffer, 2 mL 0.5 M EDTA (ethylene diamine
tetra-acetic acid), 1 mL 5M NaCl, and 20 mL of 10%
SDS (sodium dodecyl sulfate) solution to 76 mL of deionized water. For the latter, add 20 mg Proteinase-K
to 1 mL de-ionized water and store at 20°C until
use. Next remove the body from the shell. This is
fairly easy in recently thawed or relaxed specimens,
but for nonrelaxed specimens the shell may have to be
gently cracked and removed from the body. Dissect
away the head, and rinse with water. Place the tissue
in a 1.5 mL microcentrifuge tube with 500 L of NET
buffer and 10 L of Proteinase-K. Cap the tube and
place it on a 37°C mixing table for 3 h (for fixed
specimens) or 4 days (for dried specimens). The buffer
and enzyme may need daily replacement. Observe the
radula under a dissecting scope to determine if tissue
digestion is complete, and rinse it with de-ionized water several times. The supernatant can be used for extraction of DNA with phenol / chloroform techniques
(see Sambrook et al., 1989). Store the radula in 25%
ethanol until ready for mounting on a microscope
slide.
A. Taxonomic Key to Families and Selected Genera of Freshwater Gastropods
1a.
Shell an uncoiled cone (limpet, or cap shaped) (Fig. 2) .......................................................................................................................2
1b.
Shell coiled .........................................................................................................................................................................................8
2a(1a).
Apex nearly central; adult up to 12 mm; Pacific drainage. . . Family Lymnaeidae...............................................................................3
2b.
Apex to the right or left of the median line; adult 7 mm or less; dextral or sinistral ...........................................................................4
3a(2a).
Apex central (Fig. 13A) ................................................................................................................................................................Lanx
3b.
Apex anterior; Columbia river (Fig. 13B) ...............................................................................................................................Fisherola
4a(2b).
Shell dextral (apex tips to left when viewed with aperture facing up); Rocky Mountain lakes, northeastern Ontario and northcentral Quebec (Fig. 13C) .................................................Family Acroloxidae ....................................................................Acroloxus
4b.
Shell sinistral (apex tips to right when viewed with aperture facing up); throughout North America ..........Family Ancylidae............5
5a(4b).
Shell elevated, apex in midline, tinged with pink or red inside and out, radially striate, with a notch-shaped depression in unworn
specimens; apertural lip broad and flat. In southeastern rivers (Fig. 13D) ..........................................................................Rhodacmea
5b.
Shell height variable; apex in midline or to right, the same color as the rest of the shell, finely radially striate or smooth; widely distributed in lotic or lentic habitats .......................................................................................................................................................6
6a(5b).
Apex with fine radial striae, eroded in older specimens; aperture width variable, open or with a horizontal shelf in the posterior; one
lobed, flat pseudobranch; widely distributed in lotic and lentic habitats (Fig. 13E) .................................................................Ferrissia
6b.
Shell more depressed; apex without radial striae; aperture ovate to subcircular, always open; two-lobed pseudobranch, the lower
elaborately folded; in eastern states and south in lentic habitats .........................................................................................................7
7a(6b).
Apex tipped to right, tentacles colorless; In canals in southern Florida and Texas (Fig. 13F)...........................................Hebetancylus
7b.
Apex in midline; Tentacles with a black core; East of the Mississippi in lentic habitats, occasionally in south-central streams
(Fig. 13G)..............................................................................................................................................................................Laevapex
FIGURE 13 Representative limpets and planorbids. (A) The lymnaeid limpet Lanx patelloides (note large
size, west coast distribution); (B) lymnaeid limpet Fisherola nutalli; (C) Acoloxus; (D) Ancylid limpet
Rhodacmea rhodacme ( hinkezi) (note notched depression and southeastern distribution); (E) ancylid limpet
Ferrissia rivularis (note elevated shell, some species possess posterior “shelf” in shell); (F) ancylid limpet
Hebetancylus excentricus (note depressed apex to the right of midline, colorless tentacles, and southern distribution); (G) Laevapex fuscus (note obtuse apex near midline of shell, black pigmented tentacles, and widespread
distribution in eastern backwaters and southern, slow-flowing streams). (H) Armiger crista (note costae);
(I) Planorbula armigera ( jenksii) (note teeth in aperture); (J) Drepanotrema kermatoides; (K) Gyraulus
deflectus; and (L) Menetus dilatatus (A – L after Burch, 1982).
317
318
Kenneth M. Brown
8a(1b).
Shell planospiral (Fig. 2); blood (hemolymph) nearly always red (contains hemoglobin); pseudobranch (false gill) near pneumostome or anus; mantle margin simple ...............................................Family Planorbidae ..............................................................9
8b.
Shell with raised spire.......................................................................................................................................................................20
9a(8a).
Adult less than 8 mm in diameter .....................................................................................................................................................10
9b.
Adult more than 10 mm in diameter ................................................................................................................................................16
10a(9a).
Shell with transverse raised ridges (costae). Canada and northern United States. (Fig. 13H)....................................................Armiger
10b.
Shell without costae .........................................................................................................................................................................11
11a(10b).
Adults 2 mm or less in diameter; Coosa River, Alabama .................................................................................................Neoplanorbis
11b.
Adults more than 2 mm in diameter .................................................................................................................................................12
12a(11b).
Aperture or body whorl with internal “teeth” or lamellae set back one quarter whorl from aperture (Fig. 13I ...................Planorbula
12b.
Aperture or body whorl without teeth..............................................................................................................................................13
13a(12b).
Shell extremely flat, multiwhorled, or with numerous, low, close-set spiral ridges (lirae). Florida, Texas and southern Arizona
(Fig. 13J) .......................................................................................................................................................................Drepanotrema
13b.
Shell not extremely flat or multi whorled, lacking lirae ....................................................................................................................14
14a(13b).
Body whorl height equal from one side to the other (Fig. 13K) ..............................................................................................Gyraulus
14b.
Body whorl rapidly increases in height toward aperture ...................................................................................................................15
15a(14b)
Body whorl rounded (Fig. 13L) ...............................................................................................................................................Menetus
15b.
Body whorl with angular edge (carina) (Fig. 14a) ..............................................................................................................Promenetus
16a(9b).
Adult greater than 30 mm; operculum present; penis to right of mantle; gelatinous eggs; In Florida, and introduced to rivers in
central Texas....................Family Pilidae ......................................(Ampullaridae) (Fig. 18a).................................................... Marisa
16b.
Adults less than 30 mm, operculum absent ......................................................................................................................................17
17a(16b).
Shell thin, fragile, body whorl relatively depressed; Florida, Texas, Arizona (Fig. 14B) ...................................................Biomphalaria
17b.
Shell thicker, solid; body whorl often high........................................................................................................................................18
18a(17b).
Body whorl extremely large; Western in distribution (Fig. 14C) ............................................................................................Vorticifex
18b.
Body whorl not extremely large .......................................................................................................................................................19
19a(18b).
Shell spire (left side) with deep conical depression, with strong carina (Fig. 14D)..................................................................Helisoma
19b.
Shell spire (left side) with shallow depression, whorls rounded on at least one side, without carina; aperture lip sometimes flared
(Fig. 14E,F) ........................................................................................................................................................................Planorbella
20a(8b).
Shell sinistral (aperture to left when shell viewed with spire pointing away from observer); mantle margin may be digitate or
lobed..............................Family Physidae.........................................................................................................................................21
20b.
Shell dextral (aperture to right) ........................................................................................................................................................24
21a(20a)
Mantle edge with fingerlike projections ............................................................................................................................................22
21b.
Mantle edge without projections, but may be serrated .....................................................................................................................23
22a(21a).
Projections on both sides of mantle .............................................................................................................................................Physa
22b.
Projections on parietal side of mantle (Fig. 14G – I) .................................................................................................................Physella
23a(21b).
Serrations extend beyond apertural lip, partly overlap shell; Texas (Fig. 14J) .....................................................................Stenophysa
23b.
Shell elongate, surface black and glossy, sutures smooth, in wooded ponds in Canada and northern United States (Fig. 14K) .............
..................................................................................................................................................................................................Aplexa
24a(20b).
Aperture with teeth on the parietal inner margin; operculum present and calcareous, paucispiral (Fig. 2); adult length about 20 mm;
gill featherlike. Florida and southern Georgia ........................................Family Neritinidae ....................................(Fig. 14I) Neritina
24b.
Shell without teeth on parietal wall, operculum present or absent. ...................................................................................................25
FIGURE 14 Representative planorbids, physids, and neritinids. (A) Promenetus exacuous (note flaired body
whorl and carina); (B) Biomphalaria glabrata; (C) Vorticifex (Parapholyx) effusa; (D) Helisoma anceps (note
strong growth lines and carina); (E) Planorbella (Helisoma) companulata (sometimes called companulatum,
note flaired lip of aperture); (F) Planorbella (Helisoma) trivolvis (this species reaches 20 mm in diameter and is
extremely common); (G) Physella (Physa) gyrina (very similar and possibly subspecies are P. anatina and P.
virgata of the midwest and south, the only physid to reach 20 mm shell length); (H) Physella (Physa) integra
(note the more elevated spire than P. gyrina, rarely reaches 10 mm shell length); (I) Physella (Petrophysa) zionis
(note the large aperture); (J) Stenophya: (K) Aplexa elongata (note the spindle shape and lustrous almost-black
shell); and (I) Neritina reclivata (note markings on shell and teeth on parietal wall). [A – I after Burch, 1982.]
319
320
Kenneth M. Brown
25a.
Spire short, operculum multispiral; shell sometimes carinate; external, featherlike gill visible when foot is extended (Fig. 15A, B)
Family Valvatidae .....................................................................................................................................................................Valvata
25b.
Spire longer ......................................................................................................................................................................................26
26a.
Without operculum; shell relatively fragile and easily crushed; respiration by pouch in mantle cavity...............Family Lymnaeidae 27
26b.
Shell stronger; operculum and gills present..........................Subclass Prosobranchia ........................................................................33
27(26a).
With large, globose body whorl and extremely large aperture (Fig. 15C)....................................................................................Radix
27b.
Body whorl more narrow .................................................................................................................................................................28
28a(27b).
Shell very narrow and elongate; southern Ontario; north-central United States to Vermont (Fig. 15D) .....................Acella haldemani
28b.
Shell not especially narrow ...............................................................................................................................................................29
29a(28b)
Shell tear-shaped, transparent and fragile, with large, oval aperture and body whorl, small spire; amphibious. Eastern North America (Fig. 15E)................................................................................................................................................................Pseudosuccinea
29b.
Shell not extremely thin or fragile ....................................................................................................................................................30
30a(29b)
Adults more than 35 mm in length ...................................................................................................................................................31
30b.
Adults less than 35 mm in length .....................................................................................................................................................32
31a(30a).
Shell with narrow, pointed spire, moderately fragile; in marshes or streams in north-central and eastern United States, Canada
(Fig. 15F).................................................................................................................................................................Lymnaea stagnalis
31b.
Shell globose, thick, with relatively wide spire. Great Lakes and St. Lawrence River drainages (Fig. 15G)...........Bulimnea megasoma
32a(30b).
Adult shell more than 13 mm, with microscopic spiral striations or malleations; columella with a well-developed twist or plait.
Widely distributed in ponds and marshes in northern states or in alpine regions of western states (Fig. 15H) ......................Stagnicola
32b.
Adult shell less than 13 mm, spiral sculpture usually absent; columella without a twist or plait; (Fig. 15I)..............................Fossaria
33a(26b).
Operculum multispiral or paucispiral (Fig. 2), outer margins not concentric ....................................................................................34
33b.
Operculum concentric (center may be paucispiral) ...........................................................................................................................47
34a(33a).
Adult less than 7 mm; males possess verge (penis) behind right tentacle ...........................................................................................35
34b.
Adults more than 15 mm in length, males lack verge........................................................................................................................40
35a(34a).
Spire high, head – foot region divided on each side by longitudinal groove; eyes in prominent swellings on the outer bases of the tentacles; amphibious or terrestrial; crawls with steplike movement; (Fig. 16A) ..............Family Pomatiopsidae....................Pomatiopsis
35b.
Shell length variable; head – foot region not divided; eyes at same location but not on prominent swellings; totally aquatic.................
Family Hydrobiidae .........................................................................................................................................................................36
36a(35b).
Verge simple, no accessory lobes (Fig. 17A)..................................................................................................Subfamily Lithoglyphinae
................................................................................................Lepyrium, Cochliopina, Fluminicola, Antrobia, Clappia, Somatogyrus
36b.
Verge with accessory lobes (Fig. 17 B-E)...........................................................................................................................................37
37a(36b).
Verge with two arms (mitten shaped, Fig. 17B, C)............................................................................................................................38
37b.
Verge with three or more divisions (Fig. 17D, E) ..............................................................................................................................39
38a.
Verges with one small and one large lobe with glandular areas ..................................................Subfamily Nymphophilinae (Fig 17B)
.....Orygocerus, Birgella, Striobia, Marstonia, Rhapinema, Notogilla, Spilochlamys, Cincinnatia, Pyrgulopsis, Fontelicella, Natricola
38b.
Verges with branches of approximately similar diameter......................................................................................................................
Subfamily Amnicolinae (Fig. 17C) .......................................................................................Amnicola, Lyrogyrus, Hauffenia, Horatia
39a.
Verges with three accessory lobes branching off main lobe (Fig. 17D)..............................................................Subfamily Hydrobiinae
...................................................................Probythinella, Hoyia, Tryonia, Pyrgophorus, Littoridinops, Aphaostracon, Hyalopyrgus
39b.
Verge ending in three similar-sized lobes (Fig. 17E).................Subfamily Fontigentinae ........................................................Fontigens
40a(34b).
Mantle edge papillate; parthenogenetic (males absent); females brood young in a pouch posterior to head; Introduced from Florida
to Texas ...............................................................................................................Family Thiaridae .................................................41
40b.
Mantle edge smooth; dioecious (males present), females oviparous, with egg-laying sinus on the right side of the foot........................
Family Pleuroceridae ........................................................................................................................................................................42
41a(40a).
Rounded whorls with spiral grooves and transverse raised lines (costae) (Fig. 16B) ..........................Introduced in Florida to Arizona
...........................................................................................................................................................................................Melanoides
10. Mollusca: Gastropoda
FIGURE 15 Representative valvatids and lymnaeids. (A) Valvata sincera; (B) Valvata tricarinata (note
carina); (C) Radix (Lymnaea) auricularia (note expanded body whorl); (D) Acella haldemani (note extremely
narrow shell); (E) Pseudosuccinea columella (note thin, transparent shell and amphibious habit); (F) Lymnaea
stagnalis (note large and fragile shell); (G) Bulimnea megasoma (note thick, large shell); (H) Stagnicola
(Lymnaea) elodes (note that malleations sometimes present, and this species is common only in ponds and
marshes, rarely lakes); and (I) Fossaria (Lymnaea) humilis (note small size and amphibious habit). [Figures
A – I after Burch, 1982.]
321
322
Kenneth M. Brown
FIGURE 16 Representative potamiopsids, pleurocerids and bithyniids. (A) Pomatiopsis lapidaria;
(B) Melanoides tuberculata (note costae and lirae); (C) Thiara granifera (note tubercules and flattened whorls
near apex); (D) Io fluvialis (length of spines variable); (E) Leptoxis praerosa; (F) Lithasia geniculata (note
thickened anterior aperture lip and shoulders on whorls); (G) Pleurocera acuta (note acute angle on anterior
aperture and relatively flat whorls); (H) Elimia ( Goniobasis, Oxytrema) livescens (note more rounded
whorls, although there is tremendous variation in shell sculpture in this genus); and (I) Bithynia tentaculata.
[A – I after Burch, 1982.]
10. Mollusca: Gastropoda
323
FIGURE 17
Structure of the verge (penis) in various subfamilies of
hydrobiids. (A) Simple, single axis verge of subfamily Lithoglyphinae;
(B) glandular crests on verge of subfamily Nymphophilinae; (C) twoducted verge of subfamily Amnicolinae; (D) verge with accessory
lobes in subfamily Hydrobiinae; and (E) threeducted verge of subfamily Fontigentinae [A – E after Burch, 1982].
41b.
Whorls flat near spire; spiral rows of nodules on shell (Fig. 16C) .................Introduced to Florida and Texas ...........................Thiara
42a(40b).
Large, often with elongated spines and a long, anterior canal on aperture (Fig. 16D); rivers in eastern Tennessee and western
Virginia.............................................................................................................................................................................................Io
42b.
Shell size and sculpture quite variable; aperture sometimes ends with a short canal .........................................................................43
43a(42b)
Shell medium-to-small, a subglobose, or globose cone (Fig. 16E); lateral radular teeth with broad, bluntly rounded median cusps; ....
...............................................................................................................................................................................................Leptoxis
43b.
Shell usually a narrow cone; lateral radular teeth with narrow, triangular median cusps..................................................................44
44a(43b).
Shell length medium, either cone-shaped or sub-globose, often with spines or nodules; inside margin of the aperture thickened, aperture meets body whorl nearly at right angle (Fig. 16F).............................................................................................................Lithasia
44b.
Shell length and sculpture variable, base of aperture either rounded or with a short canal; inner margin of the aperture without
thickening ........................................................................................................................................................................................45
45a.
Anterior (basal tip of) aperture usually pointed; whorls flat-sided.....................Mississippi, Great Lakes and Hudson River drainages
(Fig. 16G) ............................................................................................................................................................................Pleurocera
45b.
Anterior aperture and whorls more rounded ....................................................................................................................................46
46a(45b).
In river drainages east of the Mississippi River (Fig. 16H) .........................................................................................................Elimia
46b.
In river drainages west of the Mississippi ......................................................................................................................................Juga
47a(33b).
Adults less than 15 mm; operculum calcareous; from Wisconsin to Pennsylvania and New York (Fig. 16I) ............Family Bithyniidae
Bithynia tentaculatum
47b.
Adults more than 20 mm (some up to 50 or 60 mm); operculum corneous ......................................................................................48
48a(47b).
Globose, length up to 60 mm; penis on right side of mantle; calcareous eggs. Florida ..........Family Pilidae (Ampullariidae) (Fig. 18B)
...............................................................................................................................................................................................Pomacea
48b.
Subglobose, right tentacle thicker in males, used as a penis; females ovoviviparous............Throughout the United States and Canada
Family Viviparidae ...........................................................................................................................................................................49
49a(48b).
Adults over 35 and up to 50 mm; shell relatively thin, whorls not shouldered (Fig. 18C); introduced throughout United States ..........
..................................................................................................................................................................................Cipangopaludina
49b.
Shell thick and less than 35 mm .......................................................................................................................................................50
50a(49b).
Shell sometimes with one or two spiral rows of nodules; outer margin of aperture concave (when observed from the side); inside
(columellar) margin of operculum folded inward; Alabama rivers (Fig. 18D)........................................................................Tulotoma
50b.
Shell without nodules; aperture not as above ...................................................................................................................................51
51a(50b).
Operculum with spiral center and concentric edge; whorls with a median angle or low ridge (Fig. 18E)..................................Lioplax
51b.
Operculum entirely concentric; whorls without angles or ridges ......................................................................................................52
51a(51b).
Shell often with colored bands or hirsute in juveniles; aperture nearly circular (Fig. 18F)......................................................Viviparus
51b.
Colored bands absent; aperture longer than wide, forms shoulder at junction with body whorl (Fig. 18G).......................Campeloma
324
Kenneth M. Brown
FIGURE 18
Representative ampullarids and viviparids. (A) Marisa cornuarietis (note large, planospiral
shell, occurs in southern Florida and introduced to rivers in central Texas); (B) Pomacea paludosa (large,
‘Apple’ snail common in southern Florida); (C) Cipangopaludina japonica (a large, introduced, but now common species in North America); (D) Tulotoma magnifica (in some southeastern rivers, note tubercules which
may be absent in some morphs or species); (E) Lioplax subcarinata; (F) Viviparus georgianus (common, note
circular operculum, and bands which may disappear in adults); (G) Campeloma decisum (some times referred
to as decisa; note operculum is longer than it is wide, and shouldered junction of aperture and body whorl).
[Figures A – G after Burch, 1982.]
10. Mollusca: Gastropoda
ACKNOWLEDGMENTS
I would like to thank Dr. Jeffery Tamplin for the
considerable effort involved in drafting the excellent
figures for the key, and Mr. Sean Keenan for his invaluable help in preparing the chapter.
LITERATURE CITED
Aldridge, D. W. 1982. Reproductive tactics in relation to life cycle
bioenergetics in three natural populations of the freshwater
snail, Leptoxis carinata. Ecology 63:196 – 208.
Aldridge, D. W. 1983. Physiological ecology of freshwater prosobranchs, in, Russell-Hunter, W. D. Ed., The Mollusca, Vol. 6,
pp. 329 – 358, Academic Press, Orlando, FL.
Alexander, J. E., Jr., Covich, A. P. 1991a. Predator avoidance by the
freshwater snail Physella virgata in response to the crayfish Procambarus simulans. Oecologia 87:435 – 442.
Alexander, J. E., Jr., Covich, A. P. 1991b. Predation risk and avoidance behavior by two freshwater snails. Biological Bulletin
180:387–393.
Anderson, R. M., May, R. M. 1979. Prevalence of schistosome infections within molluscan populations: observed patterns and theoretical predictions. Parasitology 79:63 – 94.
Bandel, K. 1997. Higher classification and pattern of evolution of the
gastropoda. Cour. Forsch. Inst. Senckenberg 201:57 – 81.
Barnes, R. D. 1987. Invertebrate Zoology, 5th ed., Saunders,
Philadelphia, PA.
Barnese, L. E., Lowe, R. L., Hunter, R. D. 1990. Comparative grazing efficiency of pulmonate and prosobranch snails. Journal of
the North American Benthological Society 9:35 – 44.
Baudoin, M. 1975. Host castration as a parasite strategy. Evolution
29:335 – 352.
Bieler, R. 1992. Gastropod phylogeny and systematics. Annual Review of Ecology and Systematics 23:311 – 338.
Boag, D. A. 1981. Differential depth distribution among freshwater
pulmonate snails subjected to cold temperatures. Canadian
Journal of Zoology 9:733 – 737.
Boag, D. A. 1986. Dispersal in pond snails: potential role of waterfowl. Canadian Journal of Zoology 64:904 – 909.
Boag, D. A., Pearlstone, P. S. M. 1979. On the life cycle of Lymnaea
stagnalis (Pulmonate:Gastropoda) in Southwestern Alberta.
Canadian Journal of Zoology 52:353 – 362.
Boerger, H. 1975. Movement and burrowing of Helisoma trivolvis
(Say) (Gastropoda: Planorbidae) in a small pond. Canadian
Journal of Zoology 53:456 – 464.
Bogan, A.E. 1999. North American freshwater gastropod diversity
and conservation. in: The First Symposium of the Freshwater
Mollusk Conservation Society. 17 – 19, March, 1999 Chattanooga, TN (Abstract).
Boss, K. J. 1974. Oblomovism in the Mollusca. Transactions of the
American Microscopical Society 93:460 – 481.
Bousefield, J. D. 1979. Plant extracts and chemically triggered positive rheotaxis in Biomphalaria grabrata (Say), snail intermediate
host of Schistosoma mansoni. Journal of Applied Ecology
16:681 – 690.
Bovbjerg, R. V. 1952. Ecological aspects of dispersal of the snail
Campeloma decisum. Ecology 33:169 – 176.
Bovbjerg, R. V. 1968. Responses to food in lymnaeid snails. Physiological Zoology 41:412 – 423.
Boycott, A. E. 1936. The habitats of freshwater Mollusca in Britain.
Journal of Animal Ecology 5:116 – 186.
325
Brendelberger, H. 1997. Determination of digestive enzyme kinetics:
a new method to define trophic niches in freshwater snails. Oecologia 109:34 – 40.
Brendelberger, H., Jurgens, S. 1993. Suspension feeding in Bithynia
tentaculata (Prosobranchia: Bithyniidae) as affected by body
size, food and temperature. Oecologia 94:36 – 42.
Bronmark, C. 1985a. Freshwater snail diversity: effects of pond area,
habitat heterogeneity and isolation. Oecologia 67:127 – 131.
Bronmark, C. 1985b. Interactions between macrophytes, epiphytes,
and herbivores: an experimental approach. Oikos 45:26 – 30.
Bronmark, C. 1989. Interactions between epiphytes, macrophytes
and freshwater snails: a review. Journal of Molluscan Studies
55:299 – 311.
Bronmark, C. 1994. Effects of tench and perch on interactions in a
freshwater benthic food chain. Ecology 75:1818 – 1824.
Bronmark, C., Malmqvist, B. 1986. Interactions between the leech
Glossiphonia complanata and its gastropod prey. Oecologia
69:268 – 276.
Bronmark, C., Klosiewski, S. P., Stein, R. A. 1992. Indirect effects of
predation in a freshwater, benthic food chain. Ecology 73:
1662 – 1674.
Bronmark, C., Weisner, S.E.B. 1996. Decoupling of cascading trophic
interactions in a freshwater, benthic food chain. Oecologia
108:534 – 541.
Bronmark, C., Vermaat, J. E. 1998. Chapter 3. Complex-fish-snailepiphyton interactions and their effects on submerged freshwater
macrophytes in, Jepperson, E., Sondergaard, M., Cristofferson,
K. Eds., The structuring role of submerged macrophytes in lakes,
Springer, New York, pp. 47 – 68.
Brown, D. S. 1978. Pulmonate molluscs as intermediate hosts for digenetic trematodes, in, Fretter, V., Peake, J. Eds. Pulmonates,
Vol. 2A, Systematics, evolution, and ecology. Academic Press,
New York, pp. 287 – 333.
Brown, K. M. 1979. The adaptive demography of four freshwater
pulmonate snails. Evolution 33:417 – 432.
Brown, K. M. 1982. Resource overlap and competition in pond
snails: an experimental analysis. Ecology 63:412 – 422.
Brown, K. M. 1983. Do life history tactics exist at the intraspecific
level? Data from freshwater snails. American Naturalist 121:
871 – 879.
Brown, K. M. 1985. Intraspecific life history variation in a pond
snail: The roles of population divergence and phenotypic plas
ticity. Evolution 39: 387–395.
Brown, K.M. 1997. Temporal and spatial patterns of abundance in
the gastropod assemblage of a macrophyte bed. American
Malacological Bulletin 14:27 – 33.
Brown, K. M. 1998. The role of shell strength in the foraging of crayfish for gastropod prey. Freshwater Biology 40:
255 – 260.
Brown, K. M., DeVries, D. R. 1985. Predation and the distribution
and abundance of a pulmonate pond snail. Oecologia 66:
93 – 99.
Brown, K. M., Richardson, T. D. 1988. Genetic polymorphism in
gastropods: a comparison of methods and habitat scales. American Malacological Bulletin 6:9 – 17.
Brown, K. M., Strouse, B. H. 1988. Relative vulnerability of six
freshwater gastropods to the leech Nephelopsis obscura (Verrill). Freshwater Biology 19:157 – 166.
Brown, K. M., Leathers, B. K., Minchella, D. J. 1988. Trematode
prevalence and the population dynamics of freshwater pond
snails. American Midland Naturalist 120:289 – 301.
Brown, K. M., Varza, D. E., Richardson, T. D. 1989. Life cycles and
population dynamics of two subtropical viviparid snails (Prosobranchia: Viviparidae). Journal of the North American Benthological Society 8:222 – 228.
326
Kenneth M. Brown
Brown, K.M., Lodge, D.M. 1993. The importance of specifying null
models: are invertebrates really more abundant in vegetated
habitats? Limnology and Oceanography 38: 217 – 275.
Brown, K. M., Alexander, J. E., Thorp, J. H. 1998. Differences in the
ecology and distribution of lotic pulmonate and prosobranch
gastropods. American Malacological Bulletin 14:91 – 101.
Browne, R. A. 1978. Growth, mortality, fecundity, and productivity
of four lake populations of the prosobranch snail, Viviparus
georgianus. Ecology 59: 742 – 750.
Browne, R. A. 1981. Lakes as islands: biogeographic distribution,
turnover rates, and species composition in the lakes of central
New York. Journal of Biogeography 8: 75 – 83.
Browne, R. A., Russell-Hunter, W. D. 1978. Reproductive effort in
molluscs. Oecologia 27:23 – 27.
Burch, J. B. 1989. North American Freshwater Snails. Malacological
Publications, Hamburg, MI, 365 pp.
Burky, A. J. 1971. Biomass turnover, respiration, and interpopulation
variation in the stream limpet Ferrissia rivularis. Ecological
Monographs 41: 235 – 251.
Burky, A. J. 1983. Physiological ecology of freshwater bivalves, in,
Russell-Hunter, W. D., Ed., The Mollusca, Vol. 6, Ecology, pp.
281 – 327, Academic Press, Orlando, FL.
Calow, P. 1970. Studies on the natural diet of Lymnaea peregra obtusa (Kobelt) and its possible ecological implications. Proceedings of the Malacological Society of London 39:203 – 215.
Calow, P. 1973a. Field observations and laboratory experiments on
the general food requirements of two species of freshwater
snail, Planorbis contortus Linn. and Ancylus fluviatilis. Proceedings of the Malacological Society of London 40:483 – 489.
Calow, P. 1973b. The food of Ancylus fluviatilis (Mull.), a littoral,
stone-dwelling herbivore. Oecologia 13:113 – 133.
Calow, P. 1974a. Evidence for bacterial feeding in Planorbis contortus
Linn. (Gastropoda: Pulmonata) Proceedings of the Malacological
Society of London 41:145 – 156.
Calow, P. 1974b. Some observations on the dispersion patterns of
two species populations of littoral, stone-dwelling gastropods
(Pulmonata). Freshwater Biology 4:557 – 576.
Calow, P. 1975. The respiratory strategies of two species of freshwater gastropods (Ancylus Fluviatilis Mull. and Planorbis contortus Linn.) in relation to temperature, oxygen concentration,
body size, and season. Physiological Zoology 48:114 – 129.
Calow, P. 1978. The evolution of life-cycle strategies in fresh-water
gastropods. Malacologia 17:351 – 364.
Calow, P. 1981. Adaptational aspects of growth and reproduction in
Lymnaea peregra from exposed and sheltered aquatic habitats.
Malacologia 21:5 – 13.
Calow, P. 1983. Life-cycle patterns and evolution, in, Russell-Hunter,
W. D. Ed., The Mollusca, Vol. 6, Ecology., pp. 649 – 680. Academic Press, Orlando, FL.
Calow, P., Calow, L. J. 1975. Cellulase activity and niche separation
in freshwater gastropods. Nature 255:478 – 480.
Chamberlain, N. A. 1958. Life history studies of Campeloma decisum. Nautilus 72: 22 – 29.
Cheatum, E. P. 1934. Limnological investigations on respiration, annual migratory cycle, and other related phenomena in freshwater pulmonate snails. Transactions of the American-Microscopical Society 53:348 – 407.
Clampitt, P. T. 1973. Substratum as a factor in the distribution of
pulmonate snails in Douglas Lake, Michigan. Malacologia 12:
379 – 399.
Clampitt, P. T. 1974. Seasonal migratory cycle and related movements of the fresh-water pulmonate snail, Physa integra. American Midland Naturalist 92:275 – 300.
Clarke, P. H. 1981. The freshwater molluscs of Canada. National
Museum of Natural Sciences, Ottawa, Canada.
Covich, A. P. 1977. How do crayfish respond to plants and Mollusca
as alternate food resources? Freshwater Crayfish 3:165 – 169.
Covich, A.P., Crowl, T.A., Alexander, J.E., Jr., Vaughn, C.C. 1994.
Predator-avoidance responses in freshwater decapod-gastropod
interactions mediated by chemical stimuli. Journal of the North
American Benthological Society 13:283 – 290.
Cox, L.R. 1960. Gastropoda. General characteristics of Gastropoda,
in:, Treatise on Invertebrate Paleontology, Part I. Mollusca 1,
Moore, R.C. Ed., pp. 84 – 169, Lawrence: Univ. of Kansas.
Croll, R. P. 1983. Chemoreception. Biological Reviews 58:293 – 319.
Crowl, T. A. 1990. Life-history strategies of a freshwater snail in response to stream permanence and predation: balancing conflicting demands. Oecologia 84:238 – 243.
Crowl, T. A., Covich, A. P. 1990. Predator-induced life-history shifts
in a freshwater snail. Science 247:949 – 951.
Cuker, B. E. 1983. Competition and coexistence among the grazing
snail Lymnaea, Chironomidae, and microcrustacea in an arctic
epilithic lacustrine community. Ecology 64:10 – 15.
Daldorph, P. W. G., Thomas, J. D. 1995. Factors influencing the stability of nutrient-enriched freshwater macrophyte communities:
the role of stickelbacks Pungitus pungitius and freshwater
snails. Freshwater Biology 33:271 – 289.
Davis, G. M. 1979. The origin and evolution of the Pomatiopsidae,
with emphasis on the Mekong River hydrobiid gastropods.
Monographs of the Academy of Natural Sciences, Philadelphia,
Number 20.
Davis, G. M. 1982. Historical and ecological factors in the evolution,
adaptive radiation, and biogeography of freshwater molluscs.
American Zoologist 22:375 – 395.
Dazo, B. Z. 1965. The morphology and natural history of Pleurocera
acuta and Goniobasis livescens. Malacologia 2:1 – 80.
Diamond, J. M. 1975. Assembly of species communities, in: Cody,
M., Diamond, J. M. Eds., Ecology and Evolution of Communities. pp. 342 – 344.
Dillon, R. T. 1981. Patterns in the morphology and distribution of
gastropods in Oneida Lake, New York, detected using computergenerated null hypotheses. American Naturalist 118:83 – 101.
Dillon, R. T. 1987. A new Monte Carlo method for assessing taxonomic similarity within faunal samples: reanalysis of the gastropod community of Oneida Lake, New York. American Malacological Bulletin 5:101 – 104.
Dillon, R.T., Jr., Ahlstedt, S.A. 1997. Verification of the specific status of the endangered Anthony’s river snail, Athearnia anthonyi,
using allozyme electrophoresis. Nautilus 110:97 – 101.
Dillon, R.T., Jr., Lydeard, C. 1998. Divergence among Mobile Basin
populations of the pleurocerid snail genus, Leptoxis, estimated
by allozyme electrophoresis. Malacologia 39:113 – 121.
Doremus, C. M., Harman, W. N. 1977. The effects of grazing by
physid and planorbid freshwater snails on periphyton. Nautilus
91:92 – 96.
Duch, T. M. 1976. Aspects of the feeding habits of Viviparus georgianus. Nautilus 90:7 – 10.
Duncan, C. J. 1975. Reproduction, in: Fretter and Peake, Eds., Pulmonates, Vol. 1, Functional anatomy and physiology. Academic
Press, Orlando, FL, pp. 309 – 366.
Dybdahl, M. F., Lively, C. M. 1996. The geography of coevolution:
comparative population structures for a snail and its trematode
parasite. Evolution 50:2264 – 2275.
Eckblad, J. W. 1973. Experimental predation studies of malacophagous larvae of Sepedon fuscipennis (Diptera: Sciomyzidae)
and aquatic snails. Experimental Parasitology 33:331 – 342.
Eisenberg, R. M. 1966. The regulation of density in a natural population of the pond snail, Lymnaea elodes. Ecology 47:889 – 906.
Eisenberg, R. M. 1970. The role of food in the regulation of the
pond snail, Lymnaea elodes. Ecology 51:680 – 684.
10. Mollusca: Gastropoda
Eversole, A. G. 1978. Life cycles, growth and population bioenergetics
in the snail Helisoma trivolvis (Say). Journal of Molluscan Studies 44:209 – 222.
Fretter, V.,Graham, A. 1962. British prosobranch molluscs. Ray Society, London.
Fretter, V., Peake, J. 1975. Pulmonates, Vol. 1, Functional anatomy
and physiology. Academic Press, Orlando, FL.
Fretter, V., Peake, J. 1978. Pulmonates, Vol. 2A, Systematics, Evolution, and Ecology. Academic Press, Orlando, FL.
Gerking, S. D. 1957. A method of sampling the littoral macrofauna
and its application. Ecology 38:219 – 225.
Goodrich, B. 1945. Goniobasis livescens of Michigan. Miscellaneous
Publications of the Museum of Zoology of the University of
Michigan 64:1 – 26.
Harman, W. N. 1968. Replacement of pleurocerids by Bithynia in
polluted waters of central New York. Nautilus 81:77 – 83.
Harman, W. N. 1972. Benthic substrates: their effect on fresh-water
mollusca. Ecology 53:271 – 277.
Harman, W. N., Forney, J. L. 1970. Fifty years of change in the molluscan fauna of Oneida Lake, New York. Limnology and
Oceanography 15:454 – 460.
Harman, W. N., Berg, C. O. 1971. The freshwater snails of central
New York. Search (Agriculture) 1:1 – 68.
Haynes, A., Taylor, B. J. R. 1984. Food finding and food preference
in Potamopyrgus jenkensi (E. A. Smith)(Gastropoda: Prosobranchia) Archives fur Hydrobiologie 100:479 – 491.
Hershey, A. E. 1990. Snail populations in arctic lakes: competition
mediated by predation? Oecologia 82:26 – 32.
Hershler, R. 1998. A systematic review of the hydrobiid snails (Gastropoda:Rissooidea) of the Great Basin, Western United States.
Part I. Genus Pyrgulopsis. Veliger 41:1 – 132.
Hershler, R., Frest, T.J. 1996. A review of the North American freshwater snail genus Fluminicola (Hydrobiidae). Smithsonian Contributions to Zoology. 583:1 – 41.
Hershler, R., Pierson, J.M., Krotzer, R.S. 1990. Rediscovery of Tulotoma magnifica (Conrad) (Gastropoda:Viviparidae). Proceedings of the Biological Society of Washington 103:815 – 824.
Hill, W. R., Harvey, B. C. 1990. Periphyton responses to higher
trophic levels and light in a shaded stream. Canadian Journal of
Fisheries and Aquatic Sciences 47:2307 – 2314.
Hill, W. R., Boston, H. L., Steinman, A. D. 1992. Grazers and nutrients simultaneously limit lotic primary productivity. Canadian
Journal of Fisheries and Aquatic Sciences 49:504 – 512.
Holmes, J. C. 1983. Evolutionary relationships between parasitic
helminths and their hosts. in: Futuyma, D. J., Slatkin, M. Eds.,
Coevolution, pp. 161 – 185. Sinnauer, Sunderland, MA.
Holznagel, W.E. 1998. Research note: a non-destructive method for
cleaning gastropod radulae from frozen, alcohol-fixed, or dried
material. American Malacological Bulletin 14:181 – 183.
Hunter, R. D. 1975. Growth, fecundity, and bioenergetics in three
populations of Lymnaea palustris in upstate New York. Ecology
56:50 – 63.
Hunter, R. D. 1980. Effects of grazing on the quantity and quality of
freshwater aufwuchs. Hydrobiologia 69:251 – 259.
Hunter, R. D., Russell-Hunter, W. D. 1983. Bioenergetic and community changes in intertidal aufwuchs grazed by Littorina littorea.
Ecology 64:761 – 769.
Huryn, A. D., Koebel, J. W., Benke, A. C. 1994. Life history and
longevity of the pleurocerid snail Elimia: a comparative study of
eight populations. Journal of the North American Benthological
Society 13:540 – 556.
Huryn, A. D., Benke, A. C., Ward, G. M. 1995. Direct and indirect
effects of geology on the distribution, biomass, and production
of the freshwater snail Elimia. Journal of the North American
Benthological Society 14:519 – 534.
327
Hyman, L. H. 1967. The invertebrates, Vol. VI, Mollusca I. McGraw-Hill, New York.
Jacoby, J. M. 1985. Grazing effects on periphyton by Theodoxus fluviatilis (Gastropoda) in a lowland stream. Journal of Freshwater
Biology. 3:265–274.
Jarne, P., Finet, L., Delay, B., Thaler, L. 1991. Self-fertilization versus
cross-fertilization in the hemaphroditic freshwater snail Bulinus
globosus. Evolution 45:1136 – 1146.
Jenkinson, J. J., Todd, R. M. 1998. Management of native molluscan
resources, in: Benz, G. W., Collins, D. E. Eds., Aquatic fauna in
peril: the southeastern perspective. Southeast Aquatic Research
Institute, Special Publication 1.
Johnson, S. G. 1992. Parasite-induced parthenogenesis in a freshwater snail: stable, persistent patterns of parasitism. Oceologia
89:533 – 541.
Johnson, P. D., Brown, K. M., 1997. The role of current and light in
explaining the habitat distribution of the lotic snail Elimia semicarinata. Journal of the North American Benthological Society.
15:344 – 369.
Jokinen, E. H. 1978. The aestivation pattern of a population of Lymnaea elodes. American Midland Naturalist 100:43 – 53.
Jokinen, E. H. 1982. Cipangopaludina chinensis (Gastropods:Viviparidae) in North America, review and update. Nautilus
96:89 – 95.
Jokinen, E. H. 1983. The freshwater snails of Connecticut. State Geological and Natural History Survey of Connecticut, Department of Environmental Protection, Publication 109.
Jokinen, E. H. 1985. Comparative life history patterns within a littoral zone snail community. Verh. Internat. Verein. Limnol.
22:3292 – 3299.
Jokinen, E. H. 1987. Structure of freshwater snail communities:
species – area relationships and incidence categories. American
Malacological Bulletin 5:9 – 19.
Jokinen, E. H. 1991. The malacofauna of the acid and non-acid lakes
and rivers of the Adirondack mountains and surrounding lowlands, New York state, U.S.A. Verh. Internat. Verein. Limnol.
24:2940 – 2946.
Jokinen, E. H. 1992. The freshwater snails (Mollusca: Gastropoda) of
New York state. New York State Museum Bulletin 482, 112 pp.
Jokinen, E. H., Guerette, J., Kortmann, R. W. 1982. The natural history of an ovoviviparous snail, Viviparus georgianus (Lea), in
a soft-water eutrophic lake. Freshwater Invertebrate Biology
1:2 – 17.
Jung, Y. 1992. Phylogenetic relationships of some planorbid genera
(Gastropoda: Lymnophila). Malacological Review 25:73 – 102.
Kairesalo, T., Koskimies, I. 1987. Grazing by oligochaetes and snails
on epiphytes. Freshwater Biology 17:317 – 324.
Kesler, D. H. 1981. Periphyton grazing by Amnicola limosa: an
enclosure-exclosure experiment. Journal of Freshwater Ecology
1:51 – 59.
Kesler, D. H. 1983. Cellulase activity in gastropods: should it be used
in niche separation? Freshwater Invertebrate Biology 2:173 – 179.
Kesler, D. H., Jokinen, E. H., Munns, W. R., Jr. 1986. Trophic preferences and feeding morphology of two pulmonate snail species
from a small New England pond, U.S.A.Canadian Journal of
Zoology 64:2570 – 2575.
Kesler, D. H., Munns, W. R., Jr. 1989. Predation by Belostoma flumineum (Hemiptera): an important cause of mortality in freshwater snails. Journal of the North American Benthological Society 8:342 – 350.
Kosuge, S. 1966. The family Triphoridae and its systematic position.
Malacologia 4:297 – 324.
Lamberti, G. A., Ashkenas, L. R., Gregory, S. V., 1987. Effects of three
herbivores on periphyton communities in laboratory streams.
Journal of North American Benthological Society 6:92 – 104.
328
Kenneth M. Brown
Lassen, H. H. 1975. The diversity of freshwater snails in view of the
equilibrium theory of island biogeography. Oecologia 19:1 – 8.
Levri, E. P., Lively, C. M. 1996. The effects of size, reproductive condition and parasitism on foraging behavior in a freshwater snail,
Potamopyrgus antipodarum. Animal Behavior 51:891 – 901.
Lively, C. M. 1987. Evidence from a New Zealand snail for the
maintenance of sex by parasitism. Nature 328:519 – 521.
Lively, C. M. 1989. Adaptation by a parasitic trematode to local
populations of its host. Evolution 43:1663 – 1671.
Lively, C. M., Jokela, J. 1995. Spatial variation in infection by digenetic trematodes in a population of freshwater snails (Potamopyrgus antipodarum). Oecologia 103:509 – 517.
Lodge, D. M. 1985. Macrophyte – gastropod associations: observations and experiments on macrophyte choice by gastropods.
Freshwater Biology 15:695 – 708.
Lodge, D. M. 1986. Selective grazing on periphyton: a determinant
of freshwater gastropod microdistributions. Freshwater Biology
16:831 – 841.
Lodge, D. M., Kelly, P. 1985. Habitat disturbance and the stability of
freshwater gastropod populations. Oecologia 68:111 – 117.
Lodge, D. M., Brown, K. M., Klosiewski, S. P., Stein, R. A., Covich,
A. P., Leathers, B. K., Bronmark, C. 1987. Distribution of freshwater snails: spatial scale and the relative importance of
physicochemical and biotic factors. American Malacological
Bulletin 5:73 – 84.
Lodge, D. M., Kershner, M. W., Aloi, J. E., Covich, A. P. 1994. Effects of an omnivorous crayfish (Orconectes rusticus) on a
freshwater littoral food web. Ecology 75:1265 – 1281.
Lodge, D. M., Stein, R. A., Brown, K. M., Covich, A. P., Bronmark,
C., Garvey, J. E., Klosiewski, S. P. 1998. Predicting impact of
freshwater exotic species on native biodiversity: challenges in
spatial scaling. Australian Journal of Ecology 23:53 – 67.
Lowe, R. L., Hunter, R. D. 1988. Effects of grazing by Physa integra
on periphyton community structure. Journal of the North
American Benthological Society 7:29 – 36.
Lydeard, C., Mayden, R. L. 1995. A diverse and endangered aquatic
ecosystem of the Southeast United States. Conservation Biology
9:800 – 805.
Lydeard, C., Holznagel, W. E., Garner, J., Hartfield, P., Pierson, M.
1997. A molecular phylogeny of Mobile River drainage basin
pleurocerid snails (Caenogastropoda: Cerithioidea) Molecular
Phylogenetics and Evolution 7:117 – 128.
Lydeard, C., Yoder, J.H., Holznagel, W.E., Thompson, F.G., Hartfield, P. 1998. Phylogenetic utility of the 5’-half of mitochondrial 16S rDNA gene sequences for inferring relationships of
Elimia (Cerithioidea:Pleuroceridae) Malacologia 39:183 – 193.
Macan, T. T. 1950. Ecology of freshwater Mollusca in the English
Lake District. Journal of Animal Ecology 19:124 – 146.
MacArthur, R. H., Wilson, E. O., 1967. The theory of island biogeography. Princeton University Press, Princeton, NJ. USA.
Machin, J. 1975. Water relationships, in: Fretter, V., Peake, J. Eds.,
Pulmonates, Vol. 1, Functional anatomy and physiology, pp.
105 – 164. Academic Press, Orlando, FL.
Madsen, H. 1992. Food selection by freshwater snails in the Gezira
irrigation canals, Sudan. Hydrobiologia 228:203 – 217.
Marks, J. C., Lowe, R. L. 1989. The independent and interactive effects of snail grazing and nutrient enrichment on structuring periphyton communities. Hydrobiologia 185:9 – 17.
Martin, A. W. 1983. Excretion., in: Wilbur, K. M. Ed., The molluscs,
Vol. 5, Physiology, pp. 353 – 407. Academic Press, Orlando, FL.
Martin, T. H., Crowder, L. B., Dumas, C. F., Burkholder, J. M. 1992.
Indirect effects of fish on macrophytes in Bayes Mountain Lake:
Evidence for a littoral trophic cascade. Oecologia 89:476 – 481.
McCormick, P. V., Stevenson, R. J. 1989. Effects of snail grazing
on benthic algal community structure in different nutrient
environments. Journal of the North American Benthological Society 8:162 – 172.
McMahon, R. F. 1975. Growth, reproduction, and bioenergetic variation in three natural populations of a freshwater limpet
Laevapex fuscus. Proceedings of the Malacological Society of
London 41:331 – 352.
McMahon, R. F. 1983. Physiological ecology of freshwater pulmonates, in: Russell-Hunter, W. D. Ed. The mollusca, Vol. 6,
Ecology, pp. 359 – 430, Academic Press, Orlando, FL.
McMahon, R. F., Hunter, R. D., Russell-Hunter, W. D. 1974. Variation
in aufwuchs at six freshwater habitats in terms of carbon biomass
and of carbon: nitrogen ratio. Hydrobiologia 45:391 – 404.
Merrick, G. W., Hershey, A. E., McDonald, M. E. 1991. Lake trout
(Salvelinus namaycush) control of snail density and size distribution in an arctic lake. Canadian Journal of Fisheries and
Aquatic Sciences. 48:498 – 502.
Minchella, D. J., Loverde, P. T. 1981. A cost of increased early reproductive effort in the snail Biomphalaria glabrata. American
Naturalist 118:876 – 881.
Minchella, D. J., Leathers, B. K., Brown, K. M., McNair, J. K. 1985.
Host and parasite counter-adaptations: An example from a
freshwater snail. American Naturalist 126:843 – 854.
Morton, J. E. 1955. The evolution of the Ellobiidae with a discussion
on the origin of the pulmonata. Proceedings of the Zoological
Society of London 125:127 – 168.
Mulholland, P. J., Newbold, J. D., Elwood, J. W., Horn, C. L. 1983.
The effect of grazing intensity on phosphorus spiralling in autotrophic streams. Oecologia 58:358 – 366.
Mulholland, P. J., Steinman, A. D., Palumbo, A. V., Elwood, J. W.,
Kirschtel, D. B. 1991. Role of nutrient cycling and herbivory in
regulating periphyton communities in laboratory streams. Ecology 72:966 – 982.
Neves, R. J., Bogan, A. E., Williams, J. D., Ahlstedt, S. A., Hartfield,
P. W. 1998. Status of aquatic mollusks in the southeastern
United States: A downward spiral of diversity, in: Benz, G.W.,
Collins, D. E. Eds. Aquatic fauna in peril: The southeastern
perspective. Southeast Aquatic Research Institute, Special
Publication 1.
Nystrom, P., Perez, J. R. 1998. Crayfish predation and the common
pond snail (Lymnaea stagnalis): the effect of habitat complexity
and snail size on foraging efficiency. Hydrobiologia 368:201 – 208.
Okland, J. 1983. Factors regulating the distribution of freshwater
snails (Gastropoda) in Norway. Malacologia 24:277 – 288.
Osenberg, C. W. 1989. Resource limitation, competition and the influence of life history in a freshwater snail community. Oecologia 79:512 – 519.
Pace, G. L., Szuch, E. J. 1985. An exceptional stream population of
the banded apple snail, Viviparus georgianus, in Michigan.
Nautilus 99:48 – 53.
Pechenik, J. A. 1985. Biology of the invertebrates. Prindle, Weber &
Schmidt, Boston.
Pip, E. 1986. The ecology of freshwater gastropods in the central
Canadian region. Nautilus 100:56 – 66.
Ponder, W.F., Lindberg, D.R. 1997. Towards a phylogeny of gastropod molluscs: an analysis using morphological characters. Zoological Journal of Linnaean Society 119:83 – 265.
Reavell, P. E. 1980. A study of the diets of some British freshwater
gastropods. Journal of Conchology 30:253 – 271.
Remigio, E. A., Blair, D. 1997. Molecular systematics of the freshwater snail family Lymnaeidae (Pulmonata: Basommatophora) utilizing mitochondrial ribosomal DNA sequences. Journal of
Molluscan Studies 63:173 – 185.
Richardson, T. D., Scheiring, J. F., Brown, K. M. 1988. Secondary
production of two lotic snails (Pleuroceridae: Elimia). Journal
of the North American Benthological Society 7: 234 – 245.
10. Mollusca: Gastropoda
Richardson, T. D., Brown, K. M. 1989. Secondary production of two
subtropical viviparid prosobranchs. Journal of the North American Benthological Society 8:229 – 236.
Rosemond, A. D., Mulholland, P. J., Elwood, J. W. 1993. Top – down
and bottom – up control of stream periphyton: effects of nutrients and herbivores. Ecology 74:1264 – 1280.
Russell-Hunter, W. D. 1978. Ecology of freshwater pulmonates, in: Fretter, V., Peake, J., Eds., The pulmonates, Vol. 2A, Systematics, evolution and ecology , pp. 335–383, Academic Press, Orlando, FL.
Russell-Hunter, W. D. 1983. Ecology of freshwater pulmonates, in:
Russell-Hunter, W. D. Ed., The Mollusca, Vol. 6, Ecology. pp.
335 – 383, Academic Press, Orlando, FL.
Russell-Hunter, W. D., Buckley, D. E. 1983. Actuarial bioenergetics
of nonmarine molluscan productivity. in: Russell-Hunter, W. D.,
Ed., The Mollusca, Vol. 6, Ecology. pp. 463 – 503, Academic
Press, Orlando, FL.
Saffran, K. A., Barton, D. R. 1993. Trophic ecology of Orconectes
propinquus (Girard) in Georgian Bay (Ontario, Canada). Freshwater Crayfish 9:350 – 358.
Sambrook, J., Fritch, E.F., Maniatis, T. 1989. Molecular cloning, a
laboratory manual, 2nd ed., Cold Spring Habor Laboratory
Press, Cold Spring Harbor, NY, (3 vols).
Sapp, K. H., Esch, G. W. 1994. The effects of spatial and temporal
heterogeneity as structuring forces for parasite communities in
Helisoma anceps and Physa gyrina. American Midland Naturalist 132:91 – 103.
Savino, J. F., Stein, R. A. 1982. Predator-prey interaction between
large-mouth bass and bluegills as influenced by simulated submerged vegetation. Transactions of the American Fisheries Society 111:255 – 266.
Sheldon, S. P. 1987. The effects of herbivorous snails on submerged
macrophyte communities in Minnesota lakes. Ecology 68:
1920 – 1931.
Snyder, S. C., Esch, G. W. 1993. Trematode community structure in
the pulmonate snail Physa gyrina. Journal of Parisitology
79:205 – 215.
Stein, R. A., Goodman, C. G., Marschall, E. A. 1984. Using time and
energetic measures of cost in estimating prey value for fish
predators. Ecology 65:702 – 715.
Steinman, A. D., McIntire, C. D., Lowry, R. R. 1987. Effects of herbivore type and density on chemical composition of algal assemblages in laboratory streams. Journal of North American
Benthological Society 6:189 – 197.
Swamikannu, X., Hoagland, K. D. 1989. Effects of snail grazing on
the diversity and structure of a periphyton community in an eutrophic pond. Canadian Journal of Fisheries & Aquatic Sciences
46:1698 – 1704.
Tashiro, J. S. 1982. Grazing in Bithynia tentaculata: age specific
bioenergetic patterns in reproductive partitioning of ingested carbon and nitrogen. American Midland Naturalist 107: 133 – 150.
329
Tashiro, J. S., Colman, S. D. 1982. Filter feeding in the freshwater
prosobranch snail Bithynia tentaculata: bioenergetic partitioning of ingested carbon and nitrogen. American Midland Naturalist 107:114 – 132.
Thomas, D. L., McClintock, J. B. 1990. Embryogenesis and the effects of temperature on embryonic development, juvenile
growth rates, and the onset of oviposition in the freshwater pulmonate gastropod Physella cubensis. Invertebrate Reproduction
and Development 17:65 – 71.
Thompson, F. G. 1984. The freshwater snails of Florida: A manual
for identification. Univ. of Florida Press, Gainesville, FL.
Townsend, C. R. 1975. Strategic aspects of time allocation in the ecology of a freshwater pulmonate snail. Oecologia 19:105 – 115.
Townsend, C. R., McCarthy, T. K. 1980. On the defense strategy of
Physa fontinalis (L.), a freshwater pulmonate snail. Oecologia
46:75 – 79.
Tuchman, N. C., Stevenson, P. J. 1991. Effects of selective grazing by
snails on benthic algal succession. Journal of North American
Benthological Society 10:430 – 443.
Turner, A. M. 1997. Contrasting short-term and long-term effects of
predation risk on consumer-habitat use and resources. Behavioral Ecology 8:120 – 125.
Underwood, G. J. C. 1991. Growth enhancement of the macrophyte
Ceratophylum demersum by the presence of the snail Planorbus
planorbus: the effects of grazing and chemical conditioning.
Freshwater Biology 26:325 – 334.
Vail, V. A. 1978. Seasonal reproductive patterns in three viviparid
gastropods. Malacologia 17:73 – 97.
Van Cleave, H. J., Altringer, D. A. 1937. Studies on the life cycle of
Campeloma rufrum, a freshwater snail. American Naturalist
71:167 – 184.
Van Cleave, H. J., Lederer, L. G. 1932. Studies on the life cycle of the
snail, Viviparus contectoides. Journal of Morphology 53:
499 – 522.
Vermeij, G. J., Covich, A. P. 1978. Co-evolution of freshwater
gastropods and their predators. American Naturalist 112:
833 – 843.
Weber, L. M., Lodge, D. M. 1990. Periphytic food and predatory
crayfish: relative roles in determining snail distribution. Oecologia 82:33 – 39.
Wethington, A. R., Dillon, R. T. Jr. 1993. Selfing, outcrossing, and
mixed mating in the freshwater snail Physa heterostropha: lifetime fitness and inbreeding depression. Invertebrate Biology
116: 192 – 199.
Wethington, A. R., Dillion, R. T., Jr. 1996. Gender choice and
gender conflict in a non-reciprocally mating simultaneous
hermaphrodite, the freshwater snail, Physa. Animal Behavior
51: 1107 – 1118.
Wilbur, K. M. 1983. The Mollusca, Vol. 1 – 6. Academic Press,
Orlando, FL.
This Page Intentionally Left Blank
11
MOLLUSCA: BIVALVIA
Robert F. McMahon
Arthur E. Bogan
Department of Biology
Box 19498
The University of Texas at Arlington
Arlington, TX 76019
North Carolina State Museum
of Natural Sciences
Research Laboratory
4301 Ready Creek Road
Raleigh, NC 27607
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Organ-System Function
C. Environmental and Comparative
Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Collecting, Preparation for Identification,
and Rearing
I. INTRODUCTION
North American (NA) freshwater bivalve molluscs
(class Bivalvia) fall in the subclasses Paleoheterodonta (Superfamily Unionoidea) and Heterodonta (Superfamilies
Corbiculoidea and Dreissenoidea). They have enlarged
gills with elongated, ciliated filaments for suspension feeding on plankton, algae, bacteria, and microdetritus. The
mantle tissue underlying and secreting the shell forms a
pair of lateral, dorsally connected lobes. Mantle and shell
are both single entities. During development, the right and
left mantle lobes extend ventrally from the dorsal visceral
mass to enfold the body. Each lobe secrets a calcareous
shell valve which remains connected by a mid-dorsal isthmus (Allen, 1985). Like all molluscs, the shell valves consist of outer proteinaceous and inner crystalline calcium
carbonate elements (Wilbur and Saleuddin, 1983). The lateral mantle lobes secrete shell material marked by a high
proportion of crystalline calcium carbonate making them
thick, strong and inflexible, while the mantle isthmus secretes primarily protein, forming a dorsal elastic hinge ligEcology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
A. Collecting
B. Preparation for Identification
C. Rearing Freshwater Bivalves
V. Identification of the Freshwater Bivalves
of North America
A. Taxonomic Key to the Superfamilies of
Freshwater Bivalvia
B. Taxonomic Key to Genera of
Freshwater Corbiculacea
C. Taxonomic Key to the Genera of
Freshwater Unionoidea
Literature Cited
ament uniting the calcareous valves (Fig. 1). The hinge ligament is external in all freshwater bivalves. Its elasticity
opens the valves while the anterior and posterior shell adductor muscles (Fig. 2) run between the valves and close
them in opposition to the hinge ligament which opens
them on adductor muscle relaxation.
The mantle lobes and shell completely enclose the
bivalve body, resulting in cephalic sensory structures becoming vestigial or lost. Instead, external sensory structures are concentrated on the mantle margins where they
are exposed to the external environment when the valves
open. Compared to other molluscs, the bivalve body is
laterally compressed and dorso-ventrally expanded,
adapting them for burrowing in sediments, enclosure by
shell valves and mantle protecting their soft tissues from
abrasion and preventing fine sediments from entering
the mantle cavity where they could interfere with gill
suspension feeding. These adaptations along with a
highly protrusile, muscular, spadelike foot used for burrowing, have made bivalves the most successful infaunal
suspension feeders in marine and freshwater habitats.
331
332
Robert F. McMahon and Arthur E. Bogan
FIGURE 1 General morphological features of the shells of (A) corbiculoidean, (B) unionoidean, and (C)
dreissenoidean freshwater bivalves.
The NA bivalve fauna is the most diverse in the
world, consisting of approximately 308 extent native
and seven introduced taxa (Turgeon et al., 1998). The
majority of species fall in the superfamily Unionoidea
with 278 native NA and 13 recognized subspecies in 49
genera in the family, Unionidae and five species in two
genera in the family, Margaritiferidae. Many species in
this group have unique morphological adaptations and
highly endemic, often endangered populations (Neves
et al., 1997). In the superfamily, Corbiculoidea, the
family Sphaeriidae, has 36 native and four introduced
NA species. While falling into only four genera, the
Sphaeriidae are more cosmopolitan than unionoideans
(note “unionoideans” as used here refers to all species
within the superfamily, Unionoidea while “unionid”
used later in the chapter refers only to those
unionoidean species in the family, Unionidae), several
genera and species having pandemic distributions.
Corbicula fluminea falls within the Corbiculacea (fam-
ily Corbiculidae). This species invaded NA freshwaters
in the early 1900s and now extends throughout the
coastal and southern United States and Mexico, becoming the dominant benthic species in many habitats
(McMahon, 1983a,1999). More recently, two dreissenid species, Dreissena polymorpha (zebra mussel)
and Dreissena bugensis (quagga mussel) have invaded
North America. D. polymorpha was discovered in
Lake St. Clair and Lake Erie in 1988 after introduction
in 1986. It now extends throughout the Great Lakes,
St. Lawrence River, and the Mississippi River and most
of its major tributaries (Mackie and Schloesser, 1996).
D. bugensis was first found in the Erie-Barge Canal and
eastern Lake Erie, NY, in 1991 and has since spread
through Lakes Erie and Ontario and the St. Lawrence
River (Mills et al., 1996). Both species are likely to
have been simultaneously introduced as planktonic
veliger larvae released with ballast water from ships entering the Great Lakes from ports on the Bug and
11. Mollusca: Bivalvia
333
FIGURE 2
General external anatomy of the soft tissues of (A) corbiculoidean, (B) unionoidean, and
(C) dreissenoidean freshwater bivalves.
Dnieper rivers in the Black Sea, Ukraine (Marsden
et al., 1996, Mills et al., 1996).
but their soft tissue morphologies are relatively similar
and, thus, will be discussed in general terms below.
A. External Morphology
II. ANATOMY AND PHYSIOLOGY
NA freshwater bivalves fall into three superfamilies:
Unionoidea, Corbiculoidea, and Dreissenoidea. External shell morphology among these groups vary (Fig. 1),
1. Shell
Bivalve shells consist of calcium carbonate
(CaCO3) crystals embedded in a proteinaceous matrix,
both secreted by underlying mantle tissue. In most
334
Robert F. McMahon and Arthur E. Bogan
FIGURE 3 A crosssection through the mantle and shell edges of a typical freshwater
unionoidean bivalve displaying the anatomic features of the shell, mantle, and mantle edge.
Sphaeriids have a complexed cross-lamellar shell structure and lack nacre, but their mantle
edge has a similar structure.
bivalves, the shell consists of three parts: an outer proteinaceous periostracum secreted from the periostracal
groove in the mantle edge, and underlying, calcareous
prismatic and nacre layers in which CaCO3 crystals are
embedded within an organic matrix (Fig. 3). The periostracum is initially secreted free of the other layers,
but soon fuses with the underlying prismatic layer secreted by a portion of the mantle edge just external to
the periostracal groove (Fig. 3). The prismatic layer
consists of a single layer of elongated CaCO3 crystals
oriented 90° to the horizontal shell plane (Fig. 3). The
periostracum edge seals the extrapallial space between
mantle and shell, allowing CaCO3 concentrations in
that space to reach saturation levels required for crystal
deposition (Saleuddin and Petit, 1983). The tanned
protein of the periostracum is impermeable to water,
preventing shell CaCO3 dissolution. Layers of conchiolin (i.e., tanned protein) occurring within unionoidean
shells, are suggested to retard shell dissolution in calcium poor freshwaters and are not found in the Corbiculoidea or Dreissenoidea (Kat, 1985). The nacreous
layer is continuously secreted by underlying mantle epithelium. It consists of consecutive layers of small
CaCO3 crystals deposited parallel to the shell plane
embedded in an organic matrix of chitinlike mucopolysaccharide and protein (Machado et al., 1994)
(Fig. 3). Microstructure and texture patterns of CaCO3
crystals differ among major molluscan classes and interspecifically within groups (Hedegaard and Wenk,
1998). Continual secretion of the nacreous layer thick-
ens and strengthens the shell, thus accounting for most
of its mass.
Calcium (Ca2) and bicarbonate (HCO3) necessary for shell CaCO3 crystal deposition are transported
from the external medium across the body epithelium
into the hemolymph (blood). HCO3 ions are also generated from metabolically released CO2 reacting with
hemolymph water (CO2 H2O N H HCO3).
These ions are actively transported across the mantle
into the extrapallial fluid for CaCO3 deposition into
the shell matrix (Wilbur and Saleuddin, 1983).
CaCO3 crystal formation requires release of protons (H) (Ca2 HCO3 N CaCO3 H) which
must be removed from the extrapallial fluid in order to
maintain the high pH required for CaCO3 deposition
(extrapallial fluid pH 7.4 – 8.3). It is proposed that H
combines with HCO3 to form H2CO3, followed by
dissociation into CO2 and H2O which diffuse from the
extrapallial fluid into the hemolymph. The enzyme,
carbonic anhydrase, which catalyzes this reaction, is
present in mantle tissue (Wilbur and Saleuddin, 1983).
Extrapallial fluid pH is higher in freshwater than marine bivalves, favoring shell CaCO3 deposition at the
lower Ca2 concentrations of the dilute hemolymph of
freshwater species (Wilbur and Saleuddin, 1983).
CaCO3 and shell matrix precipitate from the extrapallial fluid. Ca2 and HCO3 are actively concentrated in the extrapallial fluid, thus favoring CaCO3 deposition (Wilbur and Saleuddin, 1983). The organic
shell matrix separates individual calcareous crystals
11. Mollusca: Bivalvia
and binds them with the crystal layers into a unified
structure. It also has crystal-nucleating sites (possibly
composed of Ca2-binding polypeptides) on which
CaCO3 crystals initially form. CaCO3 crystals grow on
these sites to produce a new layer of nacre (Wilbur and
Saleuddin, 1983).
A minimum of one ATP appears to be required for
deposition of two Ca2ions, with additional ATP required for active HCO3 transport (Wilbur and Saleuddin, 1983). Thus, fast-growing bivalves or those with
massive shells must devote increased proportions of
maintenance energy to shell mineral deposition. Deposition of shell organic matrix and periostracum also requires metabolic energy in the form of four ATP for
every peptide bond. Although rarely more than 10% of
shell dry mass, the shell organic matrix can account for
30 – 50% of the total dry organic matter (i.e., shell
tissue organic matter), requiring up to one-third of the
total energy allocated to organic growth (Wilbur and
Saleuddin, 1983). Fast-growing, thin-shelled species
may devote proportionately less assimilated energy to
shell production than slower growing, thick-shelled
species, allowing greater energy allocation to tissue
growth and reproduction, thereby increasing fitness.
However, a thinner, more fragile shell increases probability of predation, lethal desiccation or damage
and / or dislodgment from the substrate, reducing fitness. Shell sculpture and shape appear highly correlated
with habitat among unionoideans. Species adapted to
soft substrates in low-flow habitats have thinner shells,
with smaller hinge teeth and greater lateral compression preventing sinking into the substrate while those
adapted to faster flowing habitats with harder substrates have, thicker, more inflated, more sculptured
shells which anchor them more firmly in the substrate,
preventing dislodgment during high water flow (Watters, 1994a). Similarly, concentric sulcations on the
shells of Corbicula fluminea increase anchoring capacity in their lotic habitats (McMahon, 1999). Therefore,
the ratio of energy allocation to shell versus tissue
growth appears to be an adaptive strategy, energetic
trade-offs between these two processes being evolved
under species-specific niche selection pressures (see Section III.B).
Shell external morphology varies among species.
Externally, shells may have concentric or radial corrugations of varying sizes and densities, ridges, pigmented
rays or blotches in the periostracum. These features,
along with shape of the posterior shell ridge and overall shell shape are major taxonomic characters for distinguishing species (see Section V). Another major external shell feature is the umbo or beak, an anteriorly
curving, dorsally projecting structure on each valve,
which is the oldest portion of the shell (Fig. 1).
335
Major internal shell features include the hinge and
projecting hinge teeth, which interlock to hold the
valves in exact juxtaposition, forming the fulcrum on
which they open and close, and various mussel scars.
These can also be major taxonomic characters (see Section V). In corbiculids, massive conical cardinal teeth
form just below the umbos (one in the right and two in
the left valve), anterior and posterior of which lie lateral teeth, usually in the form of elongate lamellae (Fig.
1A). In contrast, unionoideans have no true cardinal
teeth. Instead, massive, raised, pseudocardinal teeth develop near the umbonal end of the anterior lateral
teeth, serving a function similar to cardinal teeth. Elongated, lamellar, posterior lateral teeth extend posteriorly from the umbos (Fig. 1A). Among the
unionoideans, hinge teeth are vestigial or lost in the
tribe Anodontini. The superfamily, Dreissenoidea, is
characterized by a mussel-type shell without teeth in
which the anterior end is greatly reduced and posterior
end expanded. Thus, the umbos lie at the anterior end
making the shell decidedly, anteriorly pointed (Fig.
1C). The nacre of freshwater bivalve shells may have
species-specific colors. Internal-shell muscle scars mark
the insertion points of anterior and posterior adductor
muscles, anterior and posterior pedal (foot) retractor
muscles, pedal protractor muscles and pallial line muscles which attach the mantle margin to the shell (Figs. 2
and 3). Posteriorly, the pallial line may be indented
marking the pallial sinus into which inhalant and exhalant siphons are withdrawn during valve closure. The
extent of the pallial sinus is directly correlated with size
and length of the siphons.
2. Locomotory Structures and Burrowing
With the exception of epibenthic dreissenoids
which attach to hard substrates with proteinaceous
byssal threads, the vast majority of NA freshwater bivalves burrow in sediments. Some species lie on a hard
substrate, using the foot to wedge into crevices or under rocks. All species locomote with a muscular, motile,
anterio-ventrally directed, protrusile foot (Fig. 2).
Bivalve burrowing has been detailed by Trueman
(1983) and described for the unionidean, Anodonta
cygnea (Wu, 1987). The burrowing cycle begins with
relaxation of the adductor muscles, allowing shell
valves to open against the substratum by hinge ligament
expansion, thus anchoring the shell in place. Contraction of transverse and circular muscles around pedal hemolymph sinuses (i.e., open blood spaces) causes the
foot to narrow and lengthen, forcing it anteriorly into
the substrate. Once extended, its distal tip is anchored
either by expansion with hemolymph or lateral curving
into the substrate. Shell adductor muscles then contract,
rapidly closing the valves, releasing their hold on the
336
Robert F. McMahon and Arthur E. Bogan
substrate. Rapid valve closure expels a jet of water from
the mantle cavity, anteriorly through the pedal gape,
loosening compacted sediments at the anterior shell
margin. Thereafter, alternating contraction of anterior
and posterior pedal retractor muscles simultaneously
rock the anterior shell margin dorsoventrally and pull it
forward into the loosened sediments against the anchored foot tip. Once a new position is achieved, shell
adductor muscles relax, re-anchoring the open valves in
the substrate, reinitiating the burrowing cycle. NA
freshwater bivalves have relatively short siphons (the
mantle edges are not fused to form true siphons in
unionoideans and siphons are highly reduced or absent
in some species of Pisidium), thus, they generally are
found either just beneath the sediment surface or with
the posterior shell margins just above it. Living near the
sediment surface can lead to dislodgment, thus a
number of riverine unionoidean taxa have shell and
pedal morphologies adapted for rapid reburial (Watters,
1994a).
Many juvenile freshwater bivalves crawl considerable distances over the substrate before settlement,
holding the shell valves upright on an extended,
dorsoventrally flattened foot. Such surface locomotion
also occurs among adult sphaeriids (Wu and Trueman,
1984) and in juvenile C. fluminea (observed by the author). Crawling involves extension of the foot, anchoring its tip with mucus and / or a muscular attachment
sucker, followed by pedal retractor muscle contraction
which pulls the body forward. Pedal surface locomotion is reduced or lost in most adult unionoideans, but
is retained to varying degrees in adult sphaeriids, C.
fluminea and dreissenids. Adult dreissenids may spontaneously release from the byssus and crawl long distances before re-attachment. Single byssal threads are
produced during crawling to prevent dislodgment.
Pedal locomotion is particularly common in juvenile
and immature dreissenids and allows their escape from
locally poor conditions or highly dense mussel clumps
(Clarke and McMahon, 1996a).
B. Organ-System Function
1. Circulation
Bivalves have an ‘open’ circulatory system in which
hemolymph is not always enclosed in vessels. Rather,
circulatory fluid is carried by vessels from the heart to
various parts of the body where it passes into open,
spongy hemocoels (i.e., blood sinuses). In the hemocoels, it bathes tissues directly, percolating through
them before returning to the heart via the gills. Circulatory fluid in open systems is called “hemolymph.” Bivalves have large hemolymph volumes, accounting for
49 – 55% of total body water (Jones, 1983). The bivalve heart ventricle uniquely surrounds the rectum
and pumps oxygenated hemolymph from the gills and
mantle via the kidney through anterior and posterior
vessels (Jones, 1983, Narain and Singh, 1990) (Fig. 4)
which subdivide into smaller vessels to various parts of
the body, including pallial arteries to the mantle and
visceral arteries to the visceral mass and foot. These
secondary arteries further subdivide into many tiny vessels that open into the hemocoels where cellular exchange of nutrients, gases, and wastes occurs. Thereafter, deoxygenated hemolymph is carried from the
body tissues and organs to the mantle and gills to be
reoxygenated and, thence, to the heart. Evolution of an
open circulatory system in molluscs, including bivalves,
is associated with coelom reduction. The heart ventricle
is surrounded by a coelomic remnant, the “pericardial
cavity,” enclosed by the pericardial epithelium or “pericardium.” Other coelomic remnants include spaces
comprising the kidneys (‘coelomoducts’) and gonads
(Jones, 1983).
Like most bivalves, the hemolymph of all freshwater species has no specialized respiratory pigments for
O2 transport (Bonaventura and Bonaventura, 1983).
Instead, O2 is transported dissolved directly in the hemolymph fluid which has an O2-carrying capacity essentially equivalent to water. The very low metabolic
demands and extensive gas exchange surfaces (mantle
and gills) of bivalves allow maintenance of a primarily
aerobic metabolism in spite of a reduced hemolymph
O2-carrying capacity. As proteinaceous respiratory pigments are the primary blood pH buffer in most animals, bivalve blood acid – base balance is dependent on
other mechanisms (see Section II.C.3).
2. Gills and Gas Exchange
In lamellibranch bivalves, including all NA freshwater species, the gills are expanded beyond the requirements for gas exchange as they are also used for
suspension (filter) feeding (Fig. 2), the main mode of
food acquisition for the majority of species (see Section
III.C.2). The left and right gills, or ctenidia, consist of
an axis which extends anterolaterally along the visceral
mass. Many long, thin, inner and outer filaments extend laterally from the axis. In all NA freshwater bivalves, filaments are fused together (an evolutionarily
advanced condition) and penetrated by a series of pores
or ostia (i.e., eulamellibranch condition). From the
axis, the filaments first extend ventrally (descending filament limbs) and then reflect dorsally (ascending filament limbs) to attach distally to the dorsal mantle wall
(outer filaments) or the dorsal side of the visceral mass
(inner filaments), forming two V-shaped, porous curtains called the outer and inner “demibranchs” (Fig. 5).
11. Mollusca: Bivalvia
337
FIGURE 4
General internal anatomy, organs, and organ systems of the soft tissues of (A)
corbiculoidean, (B) unionoidean, and (C) dreissenoidean freshwater bivalves.
The demibranchs completely separate the mantle cavity
into ventral inhalant and dorsal exhalant portions. The
descending and ascending filament limbs are held
closely adjacent by tissue bridges called “interlamellar
junctions” and enclose an area called a “water tube” or
interlamellar space (Fig. 5).
Feeding and respiratory currents are sustained by
‘lateral cilia’ on the adjacent external filament surfaces
338
Robert F. McMahon and Arthur E. Bogan
FIGURE 5 The structural features of the gills (ctenidia) of freshwater bivalves. (A) Cross section through the
central visceral mass, ctenidia, and mantle of a typical freshwater unionoidean, with high magnification view
showing details of filaments, ostia, and ciliation. (B) Diagrammatic representation of the respiratory and
feeding currents across the ctenidium. (C) Diagrammatic cross-sectional representation of lammellibranch
bivalve ctenidia.
(Fig. 5). They drive water entering the inhalant mantle
cavity via the inhalant siphon or siphonal notch
through the ostia between filaments into the water
tubes. Water passes dorsally up the water tubes into
the dorsal ‘exhalant mantle cavity,’ called the
“suprabranchial cavity” or “epibranchial cavity,”
formed above the ctenidial curtain where it flows posteriorly, exiting via the exhalant siphon or siphonal
notch (Figs. 2 and 5).
While the lateral cilia drive water across the gill,
rate of water flow is partially controlled by size of the
ostia. In unionoideans, two muscle bands control ostial
pore size. Antero-posterior oriented muscle bands are
attached to vertically oriented chitinous filament supporting rods which alternate with rows of ostia. Horizontal muscle contraction pulls adjacent supporting
rods together to reduce ostial pore diameter, slowing
water flow. A second set of muscle bands run dorso-
ventrally between rows of ostia and, when contracted,
increase ostial diameter, increasing flow rate. Ostial diameter of unionoidean gills increased 2 – 3 times with
external application of the neurotransmitter, serotonin,
and external serotonin and dopamine application increased lateral cilia activity, suggesting that gill water
flow is ultimately under nervous system control (Gardiner et al., 1991). Similar muscular control of ostial
diameter occurs in Dreissena polymorpha where contraction of horizontally oriented muscle bands and
spincterlike ostial muscles reduce ostial diameter, slowing water flow. External application of serotonin relaxes these muscles increasing ostial diameter and gill
water flow (Medler and Silverman, 1997).
3. Excretion and Osmoregulation
Freshwater bivalves, like all freshwater animals,
have hemolymph and tissue osmotic concentrations
11. Mollusca: Bivalvia
greater than the very dilute freshwater medium, resulting in constant ion loss and water gain. Osmoregulatory problems are compounded by the extensive gill and
mantle surfaces of bivalves over which ion and water
fluxes occur (Dietz, 1985). To reduce these fluxes, freshwater bivalves have evolved the lowest hemolymph and
cell osmotic concentrations of any metazoan, being
20 – 50% of that found in most other freshwater species
(Dietz, 1985). In spite of their reduced osmotic concentration, the extensive epithelial surfaces of bivalves still
cause water and ion fluxes to be greater than those
recorded in other freshwater species, leading to urine
clearance rates of 20 – 50 ml/k per hour (Dietz, 1985).
In unionoideans, Na is taken up in exchange for
outward transport of the metabolic waste cations, H
and NH4, and, perhaps, Ca2. Chloride (Cl) ion uptake is in exchange for metabolically produced HCO3
or OH. Active Ca2 uptake also occurs (Burton,
1983). In freshwater snails, the major source of Ca2 is
ingested food (McMahon, 1983b), while the relative
roles of food and transport in the Ca2 balance of
freshwater bivalves are unknown. Sodium ion (Na)
uptake does not require presence of Cl which is indicative of separate transport systems for these ions
(Burton, 1983); however, freshwater dreissenids appear
to co-transport Na and Cl (Horohov et al., 1992).
While active transport is the major route by which
unionoideans gain ions, exchange diffusion (i.e., transport of an ion linked with diffusion of a second ion
species down its concentration gradient) accounts for
67% of Na uptake in C. fluminea. C. fluminea and
D. polymorpha have higher ion-transport rates than
unionoideans and sphaeriids, reflecting their geologically recent penetration of freshwaters (Dietz, 1985,
Horohov et al., 1992, Wilcox and Dietz, 1995). C. fluminea has a hemolymph solute concentration that is
higher (Dietz, 1985) and D. polymorpha, that is lower
than most other freshwater bivalves (Dietz et al.,
1996a). The low hemolymph ion-concentration of D.
polymorpha occurs in spite of its high rates of active
ion uptake, indicating that it is much more ion-permeable than other freshwater bivalves, perhaps due to its
recent evolutionary invasion of freshwaters (Dietz
et al., 1994, 1995, 1996a, b). In contrast, epithelial ion
permeability in C. fluminea is low (Zheng and Dietz,
1998a) and active ion uptake rate high (Zheng and Dietz, 1998b), allowing maintenance of higher hemolymph ion concentrations and tolerance greater ambient osmolarity variation than other freshwater
bivalves (Zheng and Dietz, 1998b). Interestingly, exchange diffusion may account for up to 90% of Cl
turnover in unionoideans in pond water, but when saltdepleted, active transport dominates Cluptake (Dietz,
1985). Na and Cl uptake can occur over the body
339
epithelia in unionoideans, but the majority occurs over
the gills (Dietz, 1985), with the epithelial cells of water
canals connecting gill ostia to water tubes being a major site for active ion and osmotic regulation (Kays
et al., 1990). Active ion uptake and hemolymph-ion
concentrations in unionoideans and, by inference,
other freshwater bivalves, may be regulated by neurotransmitter substances released by gill neurons (Dietz
et al., 1992).
Excess water is eliminated via coelomoducts or
kidneys. The heart auricle walls initially ultrafilter the
blood. Hydrostatic pressure generated by auricle contraction forces blood fluid, ions and small organic molecules through the auricle walls into the pericardial
cavity surrounding the heart. Filtration occurs through
podocyte cells of the pericardial gland lining the inner
auricular surface and, perhaps, through the efferent
branchial vein running from the longitudinal kidney
vein to the auricles (Martin, 1983) (Fig. 6). Only larger
hemolymph protein, lipid and carbohydrate molecules
cannot pass the pericardial gland filter. The filtrate exits
the pericardial cavity via left and right “renopericardial
openings” in the pericardial wall to enter the “reopericardial canals” leading to the left and right kidneys.
Larger organic waste molecules are actively transported
into the filtrate by the kidney. In the Unionoidea, the
rectal wall is extremely thin where surrounded by the
ventricle, suggesting that larger organic waste molecules may be transported through it directly into the
rectum in this region (Narain and Singh, 1990). Competed excretory fluid passes via “nephridiopores” into
the epibranchial cavity to be carried on exhalant water
flow out the exhalant siphon (Figs. 2 and 4A) (Martin,
1983).
While little studied in freshwater bivalves, the kidney is the presumed site of major active ion resorption
from the filtrate into the hemolymph. Dietz and Byrne
(1999) have demonstrated reabsorption of filtrate sulfate ion (SO42) in the kidney of D. polymorpha. As
kidney walls appear relatively impermeable to water,
active filtrate-ion resorption leads to formation of a
dilute excretory fluid facilitating excess water excretion
with filtrate osmolarity being 50% that of hemolymph
in the unionid, Anodonta cygnea (Martin, 1983).
Filtrate-ion reabsorption is less energetically expensive
than active-ion uptake from the freshwater medium
because ion-concentration gradients between coelomoduct fluid and hemolymph are far less than those
between hemolymph and freshwater.
Due to rapid water influx, an elevated excretory fluid
production is required in freshwater bivalves to maintain
osmotic balance, being approximately 0.03 mL/g wet
tissue per day in A. cygnea (Martin, 1983). In D. polymorpha excretory fluid production was 2 – 3 mL / g dry
340
Robert F. McMahon and Arthur E. Bogan
FIGURE 6
Cross-sectional representation of the anatomic features of the excretory system of a typical freshwater bivalve. Arrows indicate pathways for the excretion of excess water in the hemolymph
through the excretory system to be eliminated at the excretory pore (redrawn from Martin, 1983).
tissue per day (Dietz and Byrne, 1999) which, when converted to a wet tissue weight value of 0.1 – 0.15 mL / g
dry tissue weight per day (based on dry tissue weight being 0.05 wet tissue weight) is 3.3 – 5.0 times greater than
that of A. cygnea (Martin, 1983), reflecting the high epithelial osmotic permeability of D. polymorpha compared to other freshwater bivalves (Dietz et al., 1996a,
b). In spite of renal ion absorption, the excretory fluid of
freshwater bivalves has a considerably higher ion concentration than freshwater, making excretion a major avenue of ion loss. Ions lost by excretion and epithelial diffusion are recovered by active transport from the
medium across epithelial surfaces, particularly that of the
gills (Dietz, 1985, also see Section II.C.6).
The main nitrogenous waste product of freshwater
bivalves is ammonia (NH3) diffused across external
epithelia as dissolved NH3 or ammonium ion (NH4).
NH4 can also be actively transported to the external
medium. In marine bivalves, 65 – 72% of nitrogen
excretion is as ammonia / ammonium ion and 13 – 28%
as urea (Florkin and Bricteux-Gregoire, 1972), values
likely to be similar in freshwater species. Some freshwater unionoideans appear to be able to detoxify
ammonia / ammonium ions by conversion to urea and,
perhaps, amino acids and proteins (Mani et al., 1993).
4. Digestion and Assimilation
Freshwater bivalves are suspension (filter) feeders,
filtering algae, bacteria and suspended microdetrital particles from water flowing through the gill. Filtered material is transported by gill ciliary tracks to the “labial
palps” where it is sorted on corrugated ciliated surfaces
into food and nonfood particles before food particles are
carried by cilia to the mouth. Some freshwater species
may also utilize the foot to feed on sediment organic
detrital particles (Reid et al., 1992). Filter and pedal detritus feeding are described in Section III.C.2, while this
section is devoted to food digestion and assimilation.
11. Mollusca: Bivalvia
The bivalve mouth is a simple opening flanked laterally by left and right pairs of labial palps, whose ciliary tracks deliver filtered food from the gills to the
mouth as a constant stream of fine particles. After ingestion, food particles pass through a short, ciliated
esophagus where mucus secretion and ciliary action
bind them into a mucus string before entering the
stomach. The stomach, lying in the anterio-dorsal portion of the visceral mass (Fig. 4), is a complex structure
with ciliated sorting surfaces and openings to a number
of digestive organs and structures. On its ventral floor,
posterior to the opening of the midgut, is an elongated,
evaginated, blind-ending tube, the “style sac.” The distal end of the style sac secretes the “crystalline style,” a
long mucopolysaccharide rod that projects from the
style sac into the stomach. Style-sac cells secrete digestive enzymes into the style matrix and have cilia which
slowly rotate the style. Stomach and style pH ranges
from 6.0 – 6.9, acidity depending on phase of digestion
(Morton, 1983).
The rotating style tip projects against a chitinous
plate or “gastric shield” on the dorsal roof of the stomach. The gastric shield is penetrated by microvilli from
underlying epithelial cells, believed to release digestive
enzymes onto the shield surface (Morton, 1983). Style
rotation mixes stomach contents and winds the
esophageal mucus food string on to the style tip where
slow release of its embedded enzymes begins extracellular digestion. Abrasion of the style tip against the gastric shield, triturates food particles and allows their further digestion by enzymes released from the eroding
style matrix and from digestive epithelial cells underlying the shield.
After initial trituration and digestion at the gastric
shield, food particles released into the stomach may
have several fates. Particles are size-sorted. A ciliated
ridge, the “typhlosole,” running the length of the
midgut returns large particles to the stomach for further digestion and trituration and eventually selects and
passes indigestible particles to the hindgut / rectum for
egestion. Ciliated surfaces on the posterior “sorting
caecum” of the stomach direct sufficiently small food
particles into tubules of the “digestive diverticulum”
for final intracellular digestion (Fig. 4). The diverticula
have the lowest fluid pH of the gut (Morton, 1983).
Larger food particles falling on the caecum are recycled
into the stomach for further trituration and enzymatic
digestion, thus particles may pass over its ciliated sorting surfaces several times before acceptance for intracellular digestion or rejection into the rectum to form
feces.
Digestive cells lining the lumina of the tiny, blindending, terminal tubules of the digestive diverticula
take up fine food particles by endocytosis into food
341
vacuoles for final digestion and assimilation. After
completion of intracellular digestion / assimilation, the
apical portions of the digestive cells, which contain
food vacuoles with undigested wastes and digestive enzymes, are shed into the tubule lumina as “fragmentation spherules” which are returned to the stomach on
ciliated rejection pathways. Breakdown of fragmentation spherules in the stomach releases their food vacuole contents, hypothesized to be a major source of
stomach acidity and extracellular digestive enzymes
(Morton, 1983).
Rejected indigestible matter passes through the
short hindgut into the rectum for egestion from the
anus, opening into the epibranchial cavity on the posterior face of the posterior shell adductor muscle just upstream from the exhalant siphon or opening, allowing
feces expulsion on the exhalant current. Undigested
particles are consolidated into discrete, dense, fecal pellets by mucus secreted by the hindgut / rectum, preventing their uptake on the inhalant current.
The cerebropleural and visceral ganglia release
neurohormones which influence glycogenesis (Joosse
and Geraerts, 1983). The vertebrate glycogenic hormones, insulin and adrenalin, have similar effects on
unionoideans. Insulin injected into the Indian unionid,
Lamellidens corrianus, resulted in declining hemolymph
glucose concentration and increase in pedal and digestive diverticular glycogen stores, while adrenaline injection induced glycogen store breakdown and increased
hemolymph sugar concentration (Jadhav and Lomte,
1982b). Elevated hemolymph glucose concentrations
stimulates gut epithelial cells to produce an insulin-like
substance in the unionoideans, Unio pictorum and A.
cygnea, which stimulated activity of glucose synthetase,
increasing uptake of hemolymph glucose into glycogen
stores (Joose and Geraerts, 1983). Cerebropleural ganglionic neurosecretory hormones regulate pedal and digestive diverticular accumulation and release of protein
and nonprotein stores in L. corrianus (Jadhav and
Lomte, 1983).
5. Reproductive Structures
The paired gonads of freshwater bivalves lie close
to the digestive diverticula. Among unionoideans, their
paired condition is difficult to discern. Unionoidean gonads envelop the lower portions of the intestinal tract
and sometimes extend into the proximal portions of the
foot (Fig. 4B), while those of sphaeriids, C. fluminea,
and freshwater dreissenids lie more dorsally in the visceral mass, extending on either side of the stomach, intestine and digestive diverticula (Fig. 4A, and C;
Mackie, 1984; Claudi and Mackie, 1993). A short gonoduct leading from each gonad opens into the epibranchial cavity allowing gamete release on exhalant
342
Robert F. McMahon and Arthur E. Bogan
FIGURE 7
Schematic representations of typical reproductive systems of bivalves. (A) The primitive condition in some marine bivalve
species: male or female gametes are released from the gonoduct opening into the kidney and passed externally through the excretory pore
(shown here is a primitive marine hermaphroditic bivalve; the
anatomy is essentially similar in gonochoristic species). (B) Hermaphroditic freshwater corbiculoidean bivalves (Corbicula fluminea and
sphaeriids): male and female gametes are passed through the gonoducts into the kidney duct close to the excretory pore from which gametes are shed. (C) Gonochoristic freshwater unionoideans: gametes
pass to the outside through a gonoduct and gonopore totally separate from the kidney duct and excretory pore. (After Mackie, 1984.)
currents (Fig. 4). Among unionoideans, which are generally gonochoristic except for a few hermaphroditic
species of Anodontinae and Ambleminae (Hoeh et al.,
1995), the tracts and openings of the renal and reproductive systems are entirely separate, an advanced condition (Fig. 7C). In sphaeriids and C. fluminea, which
are hermaphroditic, the gonads have distinct regions in
which either male or female acini produce sperm or
eggs. In these latter groups and the gonochoristic freshwater dreissenids, ducts carrying eggs or sperm unite
into a single gonoduct carrying gametes from each gonad into the distal end of the kidney — allowing gamete
discharge into the epibranchial cavity via the nephridial
canal and nephridiopore (Fig. 7A) — or directly into the
nephridial canal, discharging through a common pore
on a papilla in the epibranchial cavity (Fig. 7B; Mackie,
1984; Kraemer et al., 1986).
Almost all NA freshwater bivalves are ovoviviparous, brooding developing embryos in specialized gill
marsupia. Only the dreisssenids release sperm and eggs
externally, leading to development of free swimming
larvae (Nichols, 1996). In brooding species, interlamellar spaces are modified to form gill marsupia (brood
chambers). Fully formed juveniles are released from the
marsupia via the exhalant siphon in sphaeriids and C.
fluminea. In unionoideans, bivalved “glochidia larvae”
are released from either the exhalant siphon, specialized gill pores, or ruptures in the ventral margin of the
marsupial gills. Unionoidean glochidia parasitize fish
hosts before metamorphosing into free — living juveniles. Interlamellar spaces of the outer demibranch
form marsupia in unionoideans except in the subfam-
ily, Ambleminae, which form marsupia in both demibranchs (Burch, 1975b). In contrast, marsupia form in
the inner demibranchs of sphaeriids and C. fluminea.
Among species of the unionid genus, Lampsilis, marsupia develop only in the posterior portions of the outer
demibranch from which glochidia are released through
small pores directly into the inhalant siphon (for further details, see Section III.B.1).
In C. fluminea, interlamellar marsupial brooding
spaces have no specialized structures. In sphaeriids, embryos are enclosed in specialized brood chambers evaginated from gill filaments into the interlamellar space.
Among anodontid unionids, the marsupial interlamellar
space is divided by septa into separate chambers associated with each filament. Each space is further subdivided by lateral septa into a central marsupium and inner and outer water tubes transport water from the gill
ostia to the epibranchial cavity, an advanced structure
not found in other unionoidean groups. In the primitive
unionoidean families, Margartiferidae and Ambleminae, the entire outer demibranch forms the marsupium,
while among the more advanced tribes, Pleurobemini
and Lampsilini, the marsupia are limited to specific portion of the outer demibranch (Mackie, 1984).
Some unionoidean species display sexual dimorphism, generally rare in other freshwater bivalves. In
sexually dimorphic species, glochidial incubation distends the female outer marsupial demibranch (Mackie,
1984). Thus, posterior portions of the valves of female
lampsilines are inflated compared to males, accommodating the expanded posterior marsupia. Less obvious
general inflation of the female shell occurs in the anodontids (Mackie, 1984).
Monoecious unionoideans (some species Anodontinae and Ambleminae) and all sphaeriids are generally
simultaneous hermaphrodites, concurrently producing
mature eggs and sperm. C. fluminea has an unusual
pattern of producing only eggs early after maturity
(shell length ⬇6 mm) followed later by sperm production, thereafter, remaining simultaneously hermaphroditic throughout life (Kraemer and Galloway, 1986).
Bivalves lack copulatory organs. Except for dreissenids which release both sperm and eggs externally
(Nichols, 1996), freshwater bivalves release only sperm
externally which is transported on inhalant currents of
other individuals to unfertilized eggs in the gill marsupia. Among corbiculids, self-fertilization appears common in sphaeriids, occurring near the conjunction of
male and female gonoducts (Mackie, 1984), while, in
C. fluminea, embryos frequently occur in the lumina of
gametogenic follicles and gonoducts, indicative of self
fertilization within the gonad (Kraemer et al., 1986).
Self-fertilization makes hermaphroditic species highly
invasive (see Section III.B).
11. Mollusca: Bivalvia
Temperature appears to be the main reproductive
stimulus in most freshwater bivalves. Gametogenesis
and fertilization are initiated above and maintained
within critical ambient water temperature thresholds.
Other environmental factors which may influence reproduction include: neurosecretory hormones, density
dependent factors, diurnal rhythms, food availability
and parasites. Evidence for neurosecretory and
density controls has been demonstrated in sphaeriids
(Mackie, 1984). Zebra mussels spawn in response to
the neurosecretory hormone, serotonin, and presence
of algal extracts (Ram et al., 1996). Evidence of increasing activity and metabolic rate during dark
hours in unionoideans (McCorkle et al., 1979;
Englund and Heino, 1994a), C. fluminea (McCorkleShiley, 1982) and D. polymorpha (Borcherding,
1992) suggests that spawning activity and gamete /
glochidial / juvenile release rates may also display
diurnal rhythmicity.
Sperm of freshwater bivalves may have ellipsoid or
conical nuclei and acrosomes of variable complexity
(Mackie, 1984). Unionoidean sperm have rounded or
short cylindrical heads considered a primitive condition (Rocha and Azevedo, 1990; Lynn, 1994) while
that of Dreissena and Corbicula have elongate heads
(Kraemer et al., 1986; Ram et al., 1996). The sperm of
C. fluminea is uniquely biflagellate (Kraemer et al.,
1986). Sperm with elongate heads may be adapted for
swimming in gonadal and oviductal fluids more viscous than water and are associated with internal fertilization in gonadal ducts rather in marsupia (Mackie,
1984). However, the elongate-headed sperm of D.
polymorpha is an exception to this rule, because it externally fertilizes the egg. D. polymorpha has sperm
with a straight head and an oval, bulbous acrosome
while D. bugensis sperm has a curved head with a conically shaped acrosome (Denson and Wang 1994).
There appears to be specific site for sperm recognition
and entry on the vegetal pole of unionoidean eggs
(Truncilla truncata) where the vitelline coat forms a
corrugated surface surrounding a truncated cone
(Focarelli et al., 1990).
The eggs of freshwater bivalves are round and have
greater yolk volumes than marine species with planktonic larvae. Only freshwater dreissenids have relatively
small eggs (40 – 70 m) associated with their external
fertilization and small, free-swimming veliger larva,
which grows considerably in the plankton before settlement and metamorphoses to a juvenile (219 – 365 m)
(Nichols, 1996). The larger, yolky eggs of all other
species contain nourishment supporting development to
a more advanced juvenile / glochidium stage. The
Sphaeriidae, the smallest adult freshwater bivalves, produce the largest eggs, resulting in very small brood sizes
343
ranging from 6 – 24 per adult in the genus Sphaerium,
1 – 135 per adult in the genus Musculium, and 3.3 – 6.7
per adult in the genus Pisidium (Burky, 1983). Adult
Musculium partumeium, only 4.0 mm in shell length
(SL), release juveniles of 1.4 mm SL (Hornbach et al.,
1980; Way et al., 1980). In contrast, unionoideans and
C. fluminea have smaller eggs and release smaller
glochidia / juveniles (generally 0.3 mm SL) and have
much larger brood sizes (103 – 106 per adult) (Burky,
1983; McMahon, 1999). The small eggs of zebra mussels allow them to have massive fecundities of 106 per
adult female (Nichols, 1996) required for successful external fertilization and planktonic larval development.
Evolutionary implications of bivalve fecundities are discussed in Section III.B.
The bivalve egg is surrounded by a vitelline membrane that is relatively thin in sphaeriids and thicker in
unionoideans and C. fluminea (Mackie, 1984). In
unionoideans, it remains intact throughout most of embryonic development. It disintegrates during early development in sphaeriids (Heard, 1977), allowing developing embryos to absorb nutrients from brood sacs
without embryos and / or from nutrient cells lining the
interlamellar spaces of marsupial gills. In dreissenids,
the vitelline membrane is lost early to release the freeswimming trochophore larvae which metamorphoses
into a planktonic veliger (Nichols, 1996). The vitelline
membrane is also lost early to release a free-swimming
trochophore retained through development of a juvenile clam in the marsupial gills of C. fluminea (Kraemer
and Galloway, 1986).
6. Nervous System and Sense Organs
The bivalve head, entirely enclosed within the
mantle and shell valves, is not in direct contact with the
external environment. Thus, cephalic structures including sense organs have been lost, the head having only a
mouth and associated labial palps (Fig. 2). Loss of
cephalic sense organs has lead to bivalve nervous system being far less centralized than other advanced molluscan species. A pair of cerebropleural ganglia lateral
to the esophagus near the mouth, are interconnected by
dorsal, superesophageal commissures (Fig. 8). From
these extend two pairs of nerve chords. Paired dorsal
nerve chords extend posteriorly through the visceral
mass to a pair of visceral ganglia on the anterio-ventral
surface of the posterior shell adductor muscle while
paired cerebro-pedal nerve chords innervate a pair of
pedal ganglia in the foot (Ruppert and Barnes, 1994).
The pedal and cerebropedal ganglia exert motor
control over the pedal and anterior shell adductor muscles, while motor control of the siphons and posterior
shell adductor muscle is affected by the visceral ganglia. Coordination of pedal and valve movements
344
Robert F. McMahon and Arthur E. Bogan
FIGURE 8
The anatomic features of the central nervous system of a typical unionoidean freshwater
bivalve (central nervous system anatomy is essentially similar in freshwater corbiculoidean and dreissenoidean bivalves).
during burrowing and locomotion (see Section II.A.2)
resides in the cerebropedal ganglia.
Sense organs are concentrated on the mantle edge
and in siphonal tissues most directly exposed to the external environment. Mantle edge sense organs are most
concentrated on the middle sensory mantle lobe (Fig. 3).
Photoreceptor cells detect changes in light intensity associated with shadow reflexes, phototaxis and diurnal
rhythms; while tentacles and stiffened, immobile stereocilia are associated with tactile mechanoreceptor organs perceiving direct contact displacement (touch) or
vibrations. On the siphon margins, such tactile receptors prevent drawing of large particles into the mantle
cavity. When these mechanoreceptors are impinged by
large particles, the siphons are closed by sphincter muscles and / or rapidly withdraw to prevent particle entry.
Stronger mechanical stimuli to the siphons cause the
valves to be rapidly adducted, forcing water from the
siphons, ejecting any impinging material. Under intense
mantle or siphon stimulation, siphons are withdrawn
and the valves tightly closed, a common predator defense behavior in all bivalve species.
A pair of statocysts lying near or within the pedal
ganglia are enervated by commissures from the cerebropedal ganglion (Fig. 8). Statocysts are greatly reduced in sessile marine species (i.e., cemented oysters),
suggesting their importance to locomotion and burrowing in free-living freshwater species. They are lined
with ciliary mechanoreceptors responding to pressure
exerted by a calcareous statolith or series of smaller,
granular statoconia held within the statocyst vesicle
(Kraemer, 1978). As gravity sensing organs, statocysts
detect body orientation and, thus function in geotactic
and positioning responses during burrowing and pedal
locomotory behavior.
Bivalves have a pair of organs called osphradia on
the dorsal wall of epibranchial cavity, underlying the
visceral ganglion in unionoideans and C. fluminea
(Kraemer, 1981). Because they are profusely enervated,
and neuronally connected to the visceral ganglion, shell
adductor muscles and kidney, they have been considered sense organs, but their sensory function(s) are debated. In gastropods, where osphradia are located on
the incurrent side of the ctenidium, they have been considered to be mechanoreceptors, sensing suspended
particles in inhalant water flow, or chemoreceptors.
However, in bivalves, the epibranchial (exhalant) mantle cavity position of osphradia where they receive only
filtered water and their lack of extensive neuronal gill
connections appear to preclude either a mechano- or
chemoreceptor function. Rather, bivalve osphradia are
hypothesized to be light sensing organs that control
seasonal activities such as spawning, or which provide
sensory input to kidney function, and / or diurnal patterns of shell valve adduction (Kraemer, 1981).
C. Environmental and Comparative Physiology
1. Seasonal Cycles
Freshwater bivalves display seasonal physiological
responses associated with temperature and reproductive cycles. Metabolic rates vary seasonally in freshwater bivalves (for reviews see Burky, 1983; Hornbach,
1985; McMahon, 1996, 1999) with rates being
generally greatest in summer and least in winter due to
11. Mollusca: Bivalvia
345
TABLE I Seasonal Variation in the Oxygen Consumption Rates (V̇O2) of Selected Species of
Freshwater Bivalves
Species and source
Sphaerium striatinum
Hornbach et al. (1983)
Pisidium compressum
Way and Wissing (1984)
Pisidium variabile
Way and Wissing (1984)
Pisidium walkeri
Burky and Burky (1976)
Corbicula fluminea
Williams (1985)
Dreissena polymorpha
Quigley et al. (1992)
Pyganodon grandis
Huebner (1982)
Lampsilis radiata
Huebner (1982)
a
mg dry
tissue
weight
Maximum
V̇O2
l O2/h
Minimum
V̇O2
l O2/h
25 mg
7 mg
2 mg
3 mg
0.9 mg
0.05 mg
3 mg
0.9 mg
0.05 mg
1.3 mg
0.02 mg
348 mg
204 mg
60 mg
—
31.5
15.8
8.7
0.71
0.27
0.13
1.13
0.42
0.15
1.11
0.017
430.6
308.2
143.3
7.2
1.40
0.78
0.24
0.04
0.03
0.02
0.22
0.10
0.02
0.85
0.0074
34.4
31.5
25.8
2.1
10 g
5g
5g
2g
4690
2690
2090
810
400
250
170
80
Ratio
min:max
V̇O2
Q10(Acc.)a
Seasonal
temperature
range (°C)
22.5:1
20.3:1
33.6:1
17.5:1
9.0:1
6.5:1
5.14:1
4.20:1
7.5:1
1.3:1
2.3:1
12.5:1
9.8:1
5.6:1
3.6:1
4.7
4.5
5.8
—
—
—
—
—
—
1.1
1.4
3.2
2.8
2.2
2.3
2 – 22
2 – 22
2 – 22
—
—
—
—
—
—
1 – 26
1 – 26
7 – 29
7 – 29
7 – 29
6 – 21
11.7:1
10.8:1
12.3:1
10.1:1
2.7
2.6
2.8
2.6
6 – 31
6 – 31
6 – 31
6 – 31
Q10(Acc.) is the respiratory Q10 value computed from a change in the V̇O2 value recorded for individuals field acclimated to
their respective seasonal maximum and minimum temperatures.
temperature effects (Burky, 1983; Hornbach, 1985).
Annual variation in metabolic rate can be extensive
(Table I). Maximal summer O2 consumption rates
(V̇O2) may be 20 – 33 times minimal winter rates over a
seasonal range of 2 – 22°C in Sphaerium stratinum
(Hornbach et al., 1983) or as little as 1.3 times minimal winter rates in Pisidium walkeri (1 – 26°C) (Burky
and Burky, 1976) (Table I).
Immediate temperature increases induce a corresponding metabolic rate increase in ectothermic animals such as bivalves. Acute, temperature-induced
variations in metabolic rate or V̇O2 (or in any rate function) are represented by Q10 values, the factor by which
a rate function changes with a 10°C temperature increase, computed over any temperature range as:
Q10
冢 Rate
Rate 冣
2
10/[T2 T1]
1
where Rate1 is the rate at lower temperature, Rate2 the
rate at higher temperature, T1 the lower temperature
(°C), and T2 the higher temperature (°C). Q10 for metabolic rate in the majority of ectotherms is 2 – 3, essentially that of chemical reactions. Thus, Q10 values outside this range indicate active metabolic regulation,
values 1.5 suggesting active metabolic suppression
and, 3.5, active metabolic stimulation with temperature change. Freshwater bivalve Q10 values are highly
variable within and among species, ranging from 0.2 to
14.8 among 20 species of sphaeriids (Hornbach, 1985)
and from 1.2 to 3.72 among three species of sphaeriids
(Alimov, 1975). Q10 ranged from 1.5 to 4.1 for D.
polymorpha over 10 – 30°C depending on prior acclimation temperature and temperature range of determination (Alimov, 1975; McMahon, 1996). For 10°C acclimated individuals of C. fluminea, respiratory Q10
was 2.1 over 5 – 30°C (McMahon, 1979a). Among
unionoideans, in Lampsilis siliquoidea, Q10 ranged
from 1.88 to 4.98 and in Pyganodon grandis, from
1.27 to 10.35 (Huebner, 1982). While numerous environmental and physiological factors may affect the Q10
values of freshwater bivalves, no general patterns
emerge. Rather, metabolic response to temperature
appears to have evolved under species-specific microhabitat selection pressures (Hornbach, 1985).
Temperature effects could lead to massive seasonal
metabolic fluctuations; rates being suboptimal during
colder months and supraoptimal during warmer
months. Thus, many ectothermic species display metabolic temperature acclimation, involving compensatory
adjustment of metabolic rate to a new temperature
regime over periods of a few days to several weeks.
Typically, metabolic rates are adjusted upward upon
acclimation to colder temperatures and downward
upon acclimation to warmer temperatures which
346
Robert F. McMahon and Arthur E. Bogan
dampens metabolic fluctuation with seasonal temperature change, allowing year-round maintenance of near
optimal metabolic rates. For most species, such “typical” seasonal metabolic acclimation is partial, with
metabolic rates not returning to an absolute optimal
level in a new temperature regime. The degree of metabolic temperature compensation can be detected by
comparing Q10 values of V̇O2 in instantaneous response
to acute temperature change with those measured after
acclimation [Acclimation Q10 or Q10(Acc.)]. If Q10(Acc.)
approximates 1.0, metabolic temperature compensation is nearly perfect with V̇O2 regulated near an optimal level throughout the year. If Q10(Acc.) is less than 2.0
or considerably less than acute Q10, acclimation is partial, with the metabolic rate approaching, but not
reaching, the optimal level. If Q10(Acc.) is 2 – 3 or equivalent to the acute Q10, the species is incapable of temperature acclimation. If Q10(Acc.) is greater than 3.0 or
acute Q10, inverse (reverse) acclimation is displayed in
which acclimation to a colder temperature further depresses metabolic rate, and further stimulates it on acclimation to a warmer temperature.
Three of the above acclimation patterns occur in
freshwater bivalves: (1) no capacity for acclimation (i.e.,
Q10(Acc.) equivalent to acute Q10) is displayed by the
unionids P. grandis and L. radiata (Huebner, 1982) and
D. polymorpha (Quigley et al., 1992; McMahon,
1996); (2) partial acclimation, by Pisidium walkeri
where Q10(Acc.) is considerably less than acute Q10
(Burky and Burky, 1976); and (3) reverse acclimation
(i.e., Q10(Acc.) is greater than acute Q10) displayed by
Sphaerium striatinum (Hornbach et al., 1983) and C.
fluminea (Williams and McMahon, unpublished data)
(Table I). The adaptive advantage of reverse acclimation
has been questioned because it results in massive seasonal swings in metabolic rates, but among freshwater
bivalves it may reduce utilization of energy stores during nonfeeding, overwintering periods (Burky, 1983).
V̇O2 is also related to individual size or biomass in
all animals as follows:
V̇O2 aMb
where M is the individual biomass, and a and b are constants. It can be rewritten as a linear regression in which
V̇O2 and M are transformed into logarithmic values:
Log10 V̇O2 a b(Log10 M)
where a and b are the Y-intercept (i.e., V̇O2 at Log10 M
0 or M 1) and slope (i.e., increase in Log10 V̇O2 per
unit increase in Log10 M), respectively. Thus, a measures
the relative magnitude of V̇O2 and b, the rate of increase
in V̇O2 with increasing biomass. If b 1, V̇O2 increases
in direct proportion to M. If b is 1, V̇O2 increases at a
proportionately greater rate than M, and if b is 1, V̇O2
increases at a proportionately slower rate than M. Thus,
b values 1 indicate that weight-specific V̇O2 (i.e., V̇O2
per unit body mass) decreases with increasing body mass
while b 1 indicates that it increases with increasing
mass. Conventional wisdom suggests that b values range
from 0.5 to 0.8 such that weight specific V̇O2 decreases
with increasing body mass. While generally true for vertebrates, it is less characteristic of invertebrates, and particularly of molluscs, including freshwater bivalves.
Among 14 species of sphaeriids, b ranged from 0.12 to
1.45 (Hornbach, 1985). Limited data suggest that
unionoideans have more typical b values, being 0.9 for
L. radiata, and 0.77 for P. grandis (Huebner, 1982),
with a b value range of 0.71 – 0.75 being reported for a
number of unionoideans (Alimov, 1975). A b value of
0.63 is reported for D. polymorpha (Alimov, 1975). For
C. fluminea, b ranged from 1.64 – 1.66 depending on
season, temperature and individual condition (Williams
and McMahon, unpublished data). Alimov (1975) estimated an average b of 0.73 for this species. The b value
changes with season in some species (Hornbach et al.,
1983; Way and Wissing, 1984), but remains constant in
others (Burky and Burky, 1976; Huebner, 1982) and can
also vary with reproductive condition, reported to increase in some brooding adult sphaeriids (Way and
Wissing, 1982) but not in others (Burky, 1983; Hornbach, 1985). Metabolic rate may also vary with physiologic state, increasing in starved individuals of C. fluminea (Williams and McMahon, unpublished data) and
declining in Musculium partumieum during estivation or
habitat drying (Way et al., 1981).
Comparison of a values among species of Pisidium
indicated that weight-specific V̇O2 in this group (mean
a 0.399) is about 1 / 3 that of species of Musculium
(mean a 1.605) or Sphaerium (mean a 1.439)
(Hornbach, 1985). Reduced metabolic rate in pisidiids
may reflect their reduced relative gill surface areas
(Hornbach, 1985) and hypoxic interstitial suspension
feeding habitats (Lopez and Holopainen, 1987), reduced metabolic demand of profundal pisidiids perhaps
accounting for their high tolerance of hypoxia (Burky,
1983; Holopainen, 1987).
Annually, a in C. fluminea varied from 0.12 to 1.43
(mean 0.72) (Williams and McMahon, unpublished
data) while overall a for D. polymorpha was 0.140 (Alimov, 1975). The annual range in a for P. grandis was
0.13 to 1.098 (mean 0.563) and, for L. radiata,
0.403 to 1.331 (mean 0.800) (Huebner, 1982)
while Alimov (1975) reported a to range from 0.016 to
0.096 for a number of unionoideans. The elevated a of C.
fluminea and D. polymorpha relative to unionideans and
sphaeriids (range 0.399–1.605) indicates that both
species have higher V̇O2 and metabolic rates than other
freshwater bivalves. In contrast, the low a values among
unionideans reflect their relatively low metabolic rates
compared to other freshwater bivalve groups.
11. Mollusca: Bivalvia
In bivalves, shell production and tissue growth account for a large proportion of metabolic demand,
requiring greater than 20% of total metabolic expenditure in young marine blue mussels, Mytilus edulis
(Hawkins et al., 1989). Thus, unionoideans with the
slowest growth rates among freshwater bivalves (see
section III.B.2) also have the lowest metabolic rates,
while the fast-growing C. fluminea and D. polymorpha
have the highest metabolic rates (McMahon, 1996,
1999).
In some sphaeriids, V̇O2 is influenced by growth
and reproductive cycles: maximal metabolic rates occurring during periods of peak adult and brooded juvenile growth (Burky and Burky 1976; Hornbach et al.,
1983; Way et al., 1981; Way and Wissing, 1984), perhaps as a result of the elevated metabolic demands associated with tissue growth (Hawkins et al., 1989), elevated V̇O2 of brooded developmental stages, and the
energetic costs of providing maternal metabolites to
brooded juveniles (Mackie, 1984). In contrast, metabolic rates in C. fluminea (Williams and McMahon,
unpublished data) and unionoideans (Huebner, 1982)
were unaffected by embryo brooding, perhaps because
these species do not provide maternal nourishment to
brooded embryos.
347
Filtration rates also vary seasonally and with abiotic
conditions in sphaeriids. In S. striatinum (Hornbach
et al., 1984b) and M. partumeium (Burky et al., 1985a),
filtration rates decreased with increased particle concentration and decreased temperature. Maximal filtration
rates occurred during warmer summer months and
peaked during reproduction. In M. partumeium, filtration rate and V̇O2 declined in aestivating individuals
prior to summer habitat drying (Way et al., 1981; Burky,
1983). Filtration rate is directly correlated with temperature in D. polymorpha, being maximal in summer
(MacIsaac, 1996) while it appears to be relatively temperature independent in field-acclimated specimens of C.
fluminea (Long and McMahon, unpublished data).
Freshwater bivalves also display distinct seasonal
cycles in tissue biochemical content, primarily related
to reproductive cycle. Protein, glycogen and lipid contents are maximal during gonad development and gametogenesis in the freshwater unionid, Lamellidens
corrianus, and are minimal during glochidial release
(Fig. 9A); a pattern repeated in protein and lipid
contents of individual tissues (Fig. 9B – D) (Jadhav and
Lomte, 1982a). Similarly, overwintering, nonreproductive individuals of C. fluminea had twice the biomass and greater nonproteinaceous energy stores than
FIGURE 9 Seasonal variation in the protein, lipid, and glycogen contents of the wholebody and various tissues
of the freshwater unionoidean mussel, Lamellidens corrianus, relative to the reproductive cycle. All organic
contents are expressed as percentages of wet tissue weight. (A) Annual vartiation in whole-body contents of
proteins (open histgrams), glycogen (solid histograms), and lipid (cross-hatched histograms). Remaining figures
represent levels of protein (B), lipid (C) or glycogen (D) in the mantle (open histograms), digestive diverticulum
(cross-hatched histograms), gonad (solid histograms), and foot (stippled histograms). Horizontal bars at the top
of each figure represent reproductive cycles, indicating periods during which either gonads develop or glochidia
are released. Gonad development is associated with increases in organic content, and glochidial release, with decreases in organic content of the whole body and various tissues. (From the data of Jadhav and Lomte, 1982a.)
348
Robert F. McMahon and Arthur E. Bogan
summer reproductive individuals (Williams and
McMahon, 1989). Thus, reproductive effort appears to
require massive mobilization of organic energy stores
from somatic tissues to support gonad development
and gametogenesis in freshwater bivalves.
In the sphaeriids, Sphaerium corneum and Pisidium amnicum, increase in tissue glycogen content after
fall reproduction was associated with a two-to-threefold increase in anoxia tolerance of winter- over summer-conditioned individuals, the low glycogen contents
of summer-conditioned individuals apparently reducing
their capacity for anaerobic metabolism. Thus, fall accumulation of glycogen stores not only supported
spring gametogenesis, but also provided anaerobic substrate for survival of winter anoxia induced by ice
cover (Holopainen, 1987).
2. Diurnal Cycles
Freshwater bivalves display diurnal cycles of metabolic activity. Active Na uptake peaked during dark
hours in C. fluminea (McCorkle and Shiley, 1982) and
the unionid, Toxolasma texasiensis (Graves and Dietz,
1980). This rhythmicity was lost in constant light, suggesting that ion uptake rates were driven by exogenous
changes in light intensity. Such diurnal ion transport
rhythms appear closely linked to activity rhythms.
In the unioniodeans, Ligumia subrostrata, (McCorkle
et al., 1979), and European Anodonta anatina and
Unio tumidus (Englund and Heino, 1994a) valve gaping activity peaks during dark periods. Rhythmic
gaping in L. subrostrata was lost in constant light, suggesting this behavior is also driven by exogenous variation in light intensity (McCorkle et al., 1979). Similarly,
diurnal O2 consumption rate rhythms in L. subrostrata
appeared driven by light intensity, declining with increase in, and increasing with decrease in light intensity,
however, it also had an endogenous component, persisting for 14 days in constant light (McCorkle et al.,
1979). Thus, at least some freshwater bivalves may
have diurnal activity rhythms, being most active during
dark hours. Such activity rhythms may reflect diurnal
feeding and vertical migration cycles, individuals feeding at the sediment surface at night and retreating below it during daylight to avoid visual fish and bird
predators. Interestingly, diurnal valve movements do
not occur in D. polymorpha (Walz, 1978a, b), whose
byssal attachment prevents burrowing; however, others
have reported diurnal valve movement in this species
(Borcherding, 1992).
3. Other Factors Affecting Metabolic Rates
Bivalve V̇O2 can be suppressed by pollutants such
as heavy metals, ammonia, and cyanide, which degrade
metabolic functioning (Lomte and Jadhav, 1982a) lead-
ing to extirpation (Starrett, 1971; Williams et al.,
1992) or reduced growth rates (Grapentine, 1992).
Increased levels of suspended solids impaired V̇O2 and
induced starvation in three unionoidean species, indicating interference with gill respiratory and filterfeeding currents (Aldridge et al., 1987). It also suppresses V̇O2 (Alexander et al., 1994) and filtration rate
in D. polymorpha (Lei et al., 1996). Similary, high
particle concentrations depress filtration rates in C. fluminea (Way et al., 1990a). Bivalve metabolic rates may
also be density dependent. V̇O2 declined with increased
density in the unionid, Elliptio complanata. V̇O2 was
three times greater in singly held individuals relative to
groups of seven or more, suggesting release of a
pheromone suppressing metabolic rates of nearby individuals (Paterson, 1983).
Unionoideans and sphaeriids display varying degrees of respiratory regulation when subjected to progressive hypoxia (Burky, 1983). Both D. polymorpha
(McMahon, 1996) and C. fluminea (McMahon, 1979a)
are highly O2 dependent, their V̇O2 declining proportionately with declining partial pressure of O2 (PO2). Such
species are generally relatively intolerant of prolonged
hypoxia and restricted to well-oxygenated habitats. In
contrast, other freshwater bivalve species are oxygen independent and regulate V̇O2 at relatively constant levels
with progressive hypoxia until a critical V̇O2 is reached
below which V̇O2 declines proportionately with further
decline in PO2. Such species can inhabit waters periodically subjected to prolonged hypoxia. Thus, the sphaeriids, Sphaerium simile and Pisidium casertanum, from
hypoxic profundal habitats are relatively O2 independent (Burky, 1983) while the shallow temporary pond
species, Musculium partumeium, is relatively poor O2
regulator (Hornbach, 1991). The Austrailian riverine
unionoidean, Alathyria jacksoni, rarely experiencing
hypoxia, is a relatively poor O2 regulator while a periodically hypoxic, Australian, pond species, Velesunio
ambiguus, was a strong oxygen regulator (Sheldon and
Walker, 1989). Similarly, the lentic unionioidean, A.
cygnea regulated V̇O2 at a PO2 as low as 14.3 Torr (0.9%
of full air O2 staturation), maintaining near constant hemolymph O2 concentrations during progressive hypoxia
by increasing gill ventilation (Massabuau et al., 1991)
while maintaining a near-constant heart rate (Michaelidis and Anthanasiadou, 1994). The unioniodeans, Elliptio complanata and Pyganodon grandis, from a small,
Canadian, euthrophic lake, were extreme O2 regulators,
maintaining near constant V̇O2 down to 1 mg O2/L (PO2
⬇18 Torr, 11.3% of full air O2 saturation) (Fig. 10)
(Lewis, 1984). Their capacity for extreme V̇O2 regulation
is adaptive, as winter ice cover made the lake severely
hypoxic and overwintering individuals burrow deeply
into hypoxic sediments (Lewis, 1984). In contrast, the
11. Mollusca: Bivalvia
349
FIGURE 10
Respiratory responses of the freshwater unionoidean mussels, Elliptio complanata
and Pyganodon grandis to ambient O2 concentrations declining from near full-air saturation
(8 – 9 mg O2 /l, lower horizontal axis, PO2 140 – 160 torr or mg Hg, upper horizontal axis) to the
concentration at which O2 uptake ceases. Respiratory responses of four individuals of (A) E. complanata, and (B) four individuals of P. grandis. Both species maintained normal O2 uptake rates at a
PO2 as low as 15 – 20 torr (9 – 13 percent of full-air O2 saturation) suggesting elevated capacity for
O2 regulation of oxygen uptake. (Redrawn from Lewis, 1984.)
V̇O2 of tropical and semitropical species not experiencing
winter hypoxia tends to be more O2 dependent (McMahon, 1979a; Das and Venkatachari, 1984), the V̇O2 of
the subtropical species, C. fluminea, declining to very
low levels with just a 30% decline in PO2 below full air
O2 saturation levels (McMahon, 1979a).
Some freshwater bivalves are very tolerant of extreme hypoxia or even anoxia allowing survival in hypoxic conditions below the thermocline of stratified
lakes or above reducing substrates (Butts and Sparks,
1982; Belanger, 1991). Profundal sphaeriid species tolerate extreme hypoxia and anoxia throughout summer
lake stratification (Holopainen and Jonasson, 1983),
surviving 4.5 to 200 days of anoxia, depending on
season and temperature (Holopainen, 1987). The
unionoidean A. cygnea survived 22 days of anoxia
(Zs.-Nagy et al., 1982). In contrast, other freshwater
bivalves are highly intolerant of hypoxia / anoxia, including D. polymorpha and C. fluminea, restricting
them to well-oxygenated waters (Effler and Siegfried,
1994; Johnson and McMahon, 1998; Matthews and
McMahon, 1999). Byssal thread production was inhib-
ited in D. polymorpha below a PO2 of 40 Torr (25% of
full air O2 saturation) (Clarke and McMahon, 1996b).
Individual size also affects tolerance of anoxia /
hypoxia. Juveniles of the unionoidean, Elliptio complanata, are less hypoxia-tolerant than adults (Sparks
and Strayer, 1998). In contrast, anoxia and hypoxia
tolerance decreases with increased size in C. fluminea
and D. polymorpha (Johnson and McMahon, 1998,
Matthews and McMahon, 1999). The general trend for
decreased juvenile hypoxia tolerance among freshwater
bivalves may restrict some species to well-oxygenated
habitats even though the adults may be hypoxiatolerant (Sparks and Strayer, 1998).
Under anoxia, bivalves rely on anaerobic metabolic
pathways which are not those of glycolysis; but, instead, involve simultaneous catabolism of glycogen and
aspartate or other amino acids to yield the end products, alanine and succinate. Succinate can be further
degraded into volatile fatty acids such as proprionate
or acetate (Zs.-Nagy et al., 1982; de Zwaan, 1983; van
den Thillart and de Vries, 1985). Anoxic for 6 days, A.
cygnea maintained 52 – 94% of aerobic ATP levels,
350
Robert F. McMahon and Arthur E. Bogan
higher than could occur by typical glycolytic pathways,
due to its anaerobic oxidation of succinate (Zs.-Nagy
et al., 1982). These alternative anaerobic pathways are
more efficient than glycolysis (2 mol of ATP preduced
per mole of glucose catabolized), producing 4.71 – 6.43
mol of ATP per mole of glucose catabolized (de
Zwaan, 1983). Indeed, typical glycolytic pathway enzyme activities are reduced in A. cygnea during hypoxia (Michaelidis and Athanasiadou, 1994) which favors metabolite catabolism by alternative pathways
and allows greater tolerance of hypoxia / anoxia than
less efficient glycolysis. Further, the higher ATP yields
of the alternative pathways allow excretion of anaerobic endproducts, rather than retention of acidic lactate
to deleterious levels as occurs in species dependent on
glycolysis (de Zwaan, 1983).
During anerobiosis, buildup of acidic end products
in bivalve tissues and hemolymph can cause considerable tissue and hemolymph acidosis (i.e., decrease in
pH). Without respiratory pigments, freshwater bivalve
hemolymph has little inherent buffering capacity, thus
bivalves mobilize shell calcium carbonate into the hemolymph through the mantle epithelium (Machado
et al., 1990) to buffer acidosis (Heming et al., 1988;
Byrne and McMahon, 1994; Burnett, 1997). Thus,
when anaerobic, pallial fluid pH of the unionoidean
Margaritifera margaritifera remained highly constant,
but its Ca2 concentration increased (Heming et al.,
1988). Similarly, blood Ca2 levels rose eightfold in L.
subrostrata (Dietz, 1974) and nearly fivefold in C. fluminea (Byrne et al., 1991a) under anoxia induced by
emersion in a pure N2 atmosphere. A three-fold increase in hemolymph Ca2 concentration occurred in
Pyganodon grandis made anoxic by 6 days air emersion (Byrne and McMahon, 1991).
The gills of unionoideans (Steffens et al.1985) harbor extensive extracellular calcium phosphate concretions that could buffer hemolymph pH. However, their
mass increases during prolonged hypoxia and is inversely related to hemolymph pH and directly to blood
Ca2 concentration, suggesting that Ca2 released from
the shell during hypoxia is sequestered in gill concretions to prevent its loss by excretion or diffusion. On
return to normoxia, release and redeposition in the
shell of concretion Ca2 is likely to be much less energetically demanding than replacement of lost shell Ca2
from the dilute freshwater medium (Silverman et al.,
1983). The enzyme, carbonic anhydrase, is bound to
these concretions, facilitating exchange of concretion
Ca2 (uptake or release) with the hemolymph (Istin
and Girard, 1970a, b).
4. Desiccation Resistance
Body water content in freshwater bivalves can
vary greatly seasonally and during drought periods.
Thus, many species have evolved adaptations allowing tolerance of prolonged emersion and desiccation
stress (Cáceres, 1997). Freshwater bivalves may be
emersed for weeks or months during seasonal or unpredictable droughts (Byrne and McMahon, 1994).
Lack of mobility leaves individuals stranded in air as
water levels recede, while others occur in habitats that
dry completely. Unlike other freshwater invertebrates
(Cáceres, 1997), bivalves have no obvious structures
for maintenance of aerial gas exchange when emerged,
However, they have behavioral and physiological adaptations that maintain aerobic metabolism while minimizing water loss rates (Byrne and McMahon, 1994).
Survival of emersion in freshwater bivalves is correlated with capacity to control water loss. Among freshwater species, C. fluminea and D. polymorpha are
highly emersion intolerant, tolerating only 8 – 36 days
(McMahon, 1979b) or 3 – 27 days emersion (McMahon
et al., 1993), respectively, depending on temperature
and relative humidity. Dreissena bugensis is even less
emersion tolerant than D. polymorpha (Ussary and
McMahon, 1994; Ricciardi et al.,1995). In these species
and unionoideans, death occurs in most cases at a critical threshold of water loss regardless of temperature or
relative humidity; thus, rate of water loss dictates emersion tolerance times (McMahon, 1979b; McMahon
et al., 1994; Byrne and McMahon, 1994; Ricciardi
et al., 1995). In contrast, sphaeriids inhabiting temporary ponds survive emersion for several months during
summer drying (Burky, 1983). In some species, juveniles
and adults survive emersion (Collins, 1967; McKee and
Mackie, 1980), while in others, only juveniles survive
(McKee and Mackie, 1980; Way et al., 1981). Sphaeriids burrow into sediments prior to emersion (Burky,
1983) as do some unionoideans (Cáceres, 1997).
V̇O2 declines in both emersed, aestivating M. partumeium (Way et al., 1981) and emersed individuals of
C. fluminea (McMahon and Williams, 1984; Byrne
et al., 1990). Reduction in metabolic demand in
emersed individuals allows long-term maintenance on
limited energy reserves.
Freshwater bivalves have a number of mechanisms for maintaining gas exchange while emersed.
Sphaerium occidentale is emersed for several months in
its ephemeral pond habitats. In air, its V̇O2 is 20% that
of aquatic rates. Gas exchange occurs across specialized pyramidal cells extending through shell punctae
which allows continuous valve closure, minimizing water loss (Collins, 1967). In C. fluminea, aerial V̇O2 is
21% that of aquatic rates (McMahon and Williams,
1984). Emersed specimens periodically gape and expose mantle tissue margins cemented together with
mucus (Byrne et al., 1988) during which high rates of
aerial V̇O2 are sustained while no O2 uptake occurs
during valve closure (McMahon and Williams, 1984).
11. Mollusca: Bivalvia
Bursts of metabolic heat production occur during mantle edge exposure, associated with aerial oxygen consumption and apparent repayment of an O2 debt accumulated during intervening anaerobic periods of valve
closure (Byrne et al., 1990). Thus, periodic mantle margin exposure allows maintenance of aerial gas
exchange while greatly reducing the tissue surface area
exposed and duration of exposure, greatly reducing
water loss rates (Byrne and McMahon, 1994).
Frequency and duration of mantle edge exposure is
reduced in C. fluminea with increased temperature,
decreased relative humidity and increasing duration of
emersion, suggesting that increased desiccation pressure leads to a greater reliance on anaerobic metabolism to slow evaporative water loss (Byrne et al., 1988).
Emersed unionoideans periodically expose their
mantle edges (Byrne and McMahon, 1994). Emersed
specimens of Ligumia subrostata periodically expose
mantle edges and maintain aerial V̇O2 at 21 – 23% of
aquatic rate (Dietz, 1974). Periodic mantle edge exposure also occurs in emersed specimens of M. margaritifera (Heming et al., 1988), Pyganodon grandis (Byrne
and McMahon, 1991), Pyganodon grandis, Toxolasma
parvus and Uniomersus tetralasmus (Byrne and McMahon, 1994). P. grandis has a thin shell whose margins
do not completely seal when closed and a high frequency of mantle edge exposure when emersed
(25 – 90% of emersion time), resulting in rapid water
loss and poor emersion tolerance (2 – 32 days). It
avoids desiccation by rapid down-shore migration during reductions in habitat water levels (Byrne and
McMahon, 1994). More emersion-tolerant species like
T. parvus and U. tetralasmus have thicker, tightly sealing shells and their frequency and duration of mantle
edge exposure is reduced with increasing duration of
emersion and decreasing relative humidity, conserving
water as desiccation pressure increases. T. parvus continuously closes the valves in latter stages of emersion,
allowing it to survive emersion up to 145 days (Byrne
and McMahon, 1994). U. tetralasmus is found in
small, variable-level, lentic habitats and is highly emersion tolerant. In air, it occludes the siphons with viscous mucus, preventing direct atmospheric exposure of
inner mantle tissues. Its frequency and duration of
mantle edge exposure is quite low compared to other
species and it ceases mantle edge exposure in the latter
stages of emersion, making water loss rates lower than
recorded for other freshwater bivalves and allowing it
to survive emersion for up to 578 days (Byrne and
McMahon, 1994).
In air, unionoideans and C. fluminea utilize shell
Ca2 to buffer accumulating HCO3 (Byrne and
McMahon, 1994) manifested by accumulation of Ca2
and HCO3 in mantle cavity fluids of emersed M. margaritifera (Hemming et al., 1998) and hemolymph of
351
emersed C. fluminea (Byrne et al., 1991a) and P. grandis (Byrne and McMahon, 1991). Mantle edge exposure allows release of CO2 generated by metabolic and
shell-buffering processes in both C. fluminea (Byrne
et al., 1991a) and unionoideans (Byrne and McMahon,
1991).
In emersed C. fluminea there is little evidence of O2
debt payment after re-immersion, suggesting that this
and other freshwater bivalves may remain primarily
aerobic while emersed (Byrne et al., 1990).
The Unionoidea include the most emersion-tolerant NA freshwater bivalve species (for a review, see
Byrne and McMahon, 1994). Their capacity to tolerate
prolonged emersion and / or migrate vertically with
changing water levels (White, 1979) may partially account for their dominance in larger NA river drainages
subject to extensive seasonal water-level fluctuations.
Unionoidean growth, reproduction and other lifehistory phenomena may be partially driven by seasonal
water-level variation. Thus, anthropomorphic impoundment / regulation of flow rates and levels within
these drainages could be contributing to the present
decline of their unionoidean populations and species
diversity (Williams et al., 1992).
The mode of nitrogen excretion or detoxification
during emersion is unresolved for freshwater bivalves.
Ammonium ion, NH4, is the major nitrogenous excretory product of aquatic molluscs (Bishop et al., 1983).
Due to its toxicity, NH4 is generally not accumulated
in emersed molluscs even though its high solubility precludes release as ammonia gas (NH3). When emersed,
aquatic snails detoxify NH4 by conversion to urea or
uric acid, to be excreted on re-immersion. However,
most bivalves cannot convert NH4 to urea or uric acid
(Bishop et al., 1984). Without the capacity to detoxifiy
NH4, how do freshwater bivalves tolerate emersion?
Corbicula fluminea, unlike intertidal bivalves (Bishop
et al., 1983), does not catabolize amino acids during
emersion, precluding NH4 formation (Byrne et al.
1991b). Similarly, the unionideans, Lamellidens corrianus and L. marginalis, depend almost exclusively on
carbohydrate metabolism while emersed (Lomte and
Jadhav, 1982c; Sahib et al., 1983). Interestingly, some
unionoideans may be capable of converting ammonia
to urea or other protective metabolites (Summathi and
Chetty, 1990; Mani et al., 1993) which may partially
account for their extensive emersion tolerance.
Tolerance of emersion may also be associated with
physiological and biochemical alterations in emersed
individuals. Thus, individuals of Sphaerium occidentale
and Musculium securis from an emersed population
were more emersion tolerant than individuals from an
immersed, active population (McKee and Mackie,
1980), suggesting that gradual emersion may induce
emersion resistant biochemical and physiological
352
Robert F. McMahon and Arthur E. Bogan
alterations such as a shift to carbohydrate dominated
catabolism and reduced metabolic demand. Such biochemical and physiological compensation may be mediated by neurosecretory hormones (Lomte and Jadhav,
1981a).
There have been almost no studies of freeze tolerance in freshwater bivalves even though many species
occupy shallow, temperate habitats in which winter
water-level reduction could emerse individuals in subfreezing air. Marine intertidal mytilid mussels tolerate
exposure to subfreezing conditions by allowing freezing
of hemolymph and interstitial fluids while preventing
cell freezing, an adaptation manifested by their lack of
hemolymph supercooling during freezing (Aarset,
1982). In contrast, D. polymorpha is intolerant of
emersion below 3°C (Clarke et al., 1993) and displays hemolymph supercooling (Paulkstis et al., 1996).
Poor freeze tolerance in D. polymorpha may reflect its
tendency to settle at depths of 1 m, preventing winter
emersion (Clarke et al., 1993). Freeze sensitivity / tolerance in other freshwater bivalves (particularly shallow
water species) is ripe for further study.
5. Gill Calcium Phosphate Concretions
in Unionoideans
Dense calcium phosphate [Ca3(PO4)2] concretions
occur in unionoidean tissues. Mantel concretions may
provide Ca2 for shell deposition (Davis et al., 1982;
Jones and Davis, 1982; Istin and Girard, 1970a). The
mantle contains high concentrations of the enzyme,
carbonic anhydrase which may be bound to the concretions (Istin and Girard, 1970b). This enzyme catalyzes
the reaction of CO2 and H2O to form bicarbonate ion
(HCO3), suggesting that it facilitates mobilization of
concretion Ca2.
Dense extracellular calcium phosphate concretions
(diameter 1 – 3 m) also occur in unionoidean gills
(Silverman et al., 1983, 1988; Steffens et al., 1985).
They develop from amorphous material initially concentrated in membrane bound vesicles in gill connective
tissue cells (Silverman et al., 1989). Concretions account for up to 60% of gill dry weight in some species
(Silverman et al., 1985). They are most dense along
parallel nerve tracts oriented at 90° to the gill filaments
(Silverman et al., 1983; Steffens et al., 1985). They are
25% protein by dry weight. One of the proteins is similar to vertebrate, calcium-binding calmodulin. This
protein is most abundant in protein granules of concretion-forming cells prior to concretion mineral deposition, suggesting that it acts as site for calcium phosphate deposition (Silverman et al., 1988).
Besides storing shell Ca2 released to the hemolymph to buffer respiratory acidosis (see Section
II.C.3), gill concretions provide a ready source of
maternal Ca for shell deposition in brooded glochidia
(Silverman et al., 1985, 1987). Thus, gill concretion
mass in brooding individuals of L. subrostrata and P.
grandis was only 47 and 70% that of nonbrooding individuals, respectively (Silverman et al., 1985). 45Ca
tracer studies indicate that 90% of glochidial shell Ca
was of maternal origin in P. grandis, the most likely
source being gill concretions, with nonmineralized Ca
accounting for only 8% of glochidial Ca in L. subrostrata (Silverman et al., 1987).
6. Water and Salt Balance
In hypo osmotic freshwater, bivalves lose ions and
gain water from their hyperosmotic tissues and hemolymph. Excess body water is excreted as a fluid
hypo-osmotic to tissues and hemolymph via the kidneys
while ions lost in excretion and over surface epithelia
are actively recovered over the gills and epithelial tissues
as described in Section II.B.3 (reviewed by Deaton and
Greenberg, 1991). Relatively salinity tolerant C. fluminea survives exposure to a salinity of 10 – 14 ppt
(Morton and Tong, 1985), above which it remains isosmotic to the medium (Gainey and Greenberg, 1977).
Unionoideans and D. polymorpha have lower hemolymph osmolarities than C. fluminea (Dietz et al.,
1996a) and generally lose capacity for osmotic and volume regulation above 3 – 4 ppt (Hiscock, 1953; Wilcox
and Dietz, 1998). C. fluminea regulates fluid volume by
actively increasing hemolymph free amino acid concentration in hyperosmotic media (Gainey and Greenberg,
1977; Matsushima et al., 1982), preventing water loss
by active equilibration of hemolymph and medium osmolarity. Such volume regulation does not occur in
other freshwater bivalves because their low tissue osmolarities limit the free amino acid pool (Yamada and
Matsushima, 1992; Dietz et al., 1996a,b, 1998). Volume regulation may have been lost in unionoideans and
D. polymorpha, because they are not exposed to hyperosmotic conditions in freshwater. Instead, both groups
retain only limited cell volume regulation through regulation of intracellular inorganic ion concentrations
(Dietz et al., 1996a, b, 1998). The elevated osmotic
concentration and free amino acid pool volume regulation of C. fluminea reflects its recent evolution from an
estuarine ancestor (Dietz, 1985) in which such adaptations are common (Deaton and Greenberg, 1991).
Both unionoideans and C. fluminea respond to
maintenance in very dilute media by increasing active
Na uptake rate to maintain hemolymph ion concentration (Dietz, 1985). The activity of (Na and K)-activated ATPase, an enzyme required for active Na /K
transport, increased in mantle and kidney tissues of saltdepleted C. fluminea, suggesting activation of Na transport. This response did not occur in the salt-depleted
11. Mollusca: Bivalvia
Lampsilis straminca claibornesis, suggesting that active
ion uptake regulation has been lost in unionoideans with
a longer freshwater fossil history than C. fluminea
(Deaton, 1982). In C. fluminea Cl is the major blood
ion, maintained by elevated active epithelial Cl uptake.
In contrast, in the unionoidean, Toxolasma texasiensis,
Cl and HCO3 are equivalent hemolymph anionic
components. Because HCO3 can be readily mobilized
by respiratory and metabolic processes, unionoideans
may depend to a greater extent on active Na relative to
Cl uptake to maintain hemolymph osmotic balance
(Byrne and Dietz, 1997). The anionic hemolymph component of D. polymorpha, like that of C. fluminea, is
dominated by Cl, but this species actively absorbs
equal levels of Na and Cl to maintain ionic stasis, suggesting co-transport of these ions (Horohov et al.,
1992). Retention of Cl as the main hemolymph anion
in C. fluminea and D. polymorpha reflects their recent
evolution from estuarine ancestors (Deaton and Greenberg, 1991; Byrne and Dietz, 1997; Horohov et al.,
1992), while the greater dependence of C. fluminea on
Cl uptake for ionic regulation and its increased hemolymph osmotic concentration (Dietz et al., 1994) suggests that it entered freshwater more recently than D.
polymorpha. Interestingly, although D. polymorpha can
hyperosmotically regulate hemolymph ion concentrations in dilute freshwaters, it is less tolerant of elevated
medium ion concentrations than other bivalves restricting its penetration of estuarine habitats (Horohov et al.,
1992; Dietz et al., 1994, 1998; Wilcox and Dietz, 1998).
In the Asian unionoidean, Anodonta woodiana,
mantle cavity water osmolarity at 34 mosmol/L was
76% that of the hemolymph (45 mosmol/L) with pallial fluid Na, K, and Cl concentrations being 71, 76
and 72% those of the hemolymph, respectively. Maintenance of elevated mantle water ion concentrations
suggests that it acts as an osmotic buffer, reducing the
gradient for, and, thus, the rate of diffusive ion loss to
the dilute freshwater medium (Matsushima and Kado,
1982). However, loss of a hyperosmotic mantle fluid to
the external medium through the exhalant siphon
could also be a major route for ion loss.
In unionoideans and C. fluminea, the enzyme,
carbonic anhydrase (CA), which catalyzes formation of
carbonic acid (H2CO3) from water and carbon dioxide,
occurs in gill and mantle tissues (Henery and Saintsing,
1983). H2CO3 degrades into H and CO3 (bicarbonate
ion) which are counter ions for active Na and Cl uptake (Horohov et al., 1992; Dietz et al., 1994; Byrne and
Dietz, 1997). Gill and mantle CA activity increases when
freshwater bivalves are held in extremely dilute media
while CA inhibition by actazolamide results in reduced
Na and Cl uptake rates, strong evidence of the role of
CA in ion regulation (Henery and Saintsing, 1983).
353
Freshwater bivalve hemolymph osmotic concentrations are the lowest recorded for multicellular invertebrates (Dietz, 1985; Deaton and Greenberg, 1991).
Among unionoideans, hemolymph osmolarity for the
European species, Anodonta cygnea, was 40 – 50
mosmol/L or 4 – 5% that of seawater. D. polymorpha
has the lowest hemolymph concentration of all freshwater bivalves (30 – 36 mosmol/L or 3.0 – 3.6% that of
seawater) (Dietz et al., 1994), while C. fluminea has the
highest value at 65 mosmol /L (6.5% that of seawater)
(Zheng and Dietz, 1998a). That for other freshwater
invertebrates ranges from 100 to 400 mosmol/L or
10 – 40% that of seawater (Burky, 1983). Low hemolymph osmotic concentrations in freshwater bivalves reduce the gradient for transepithelial osmosis
and ion diffusion across their extensive mantle and gill
epithelial surfaces to that which can be balanced by
water excretion and active ion uptake at energetically
feasible levels (Burton, 1983). Hydrostatic pressure
generated by heart beat in A. cygena allows filtration of
enough hemolymph plasma into the pericardial space
to account for urine production and is twice that of the
marine clam, Mya arenaria, suggesting that freshwater
bivalves can excrete water at higher rates than less hypoosmotically stressed estuarine species (Jones and
Peggs, 1983). Exposure of Pyganodon sp. to a very dilute medium induced formation of extensive extracellular membrane spaces in the deep infoldings of kidney
epithelial cells, perhaps increasing surface area for active ion uptake from excretory fluids producing a more
dilute urine and / or allowing increased excretion of excess water (Khan et al., 1986).
Increasing osmotic gradients for water uptake from
marine through estuarine into freshwater habitats required that the external epithelia of bivalves became increasing impermeable to water with their evolutionary
transition through these environments. Thus, increased
epithelial osmo-resistance characterizes all freshwater
bivalves (Deaton and Greenberg, 1991; Dietz, 1985; Dietz et al., 1994; Byrne and Dietz, 1997; Zheng and Dietz, 1998a). Among freshwater bivalves, C. fluminea is
least osmotically permeable (Zheng and Dietz, 1998a)
and D. polymorpha most permeable (Dietz et al.,
1995), while unionoideans are of intermediate permeability (Dietz et al., 1996a, b; Zheng and Dietz, 1998a).
The “osmotically tight” epithelium of C. fluminea allows it to maintain higher hemolymph osmolarity than
other freshwater bivalves while producing normal excretory fluid volumes (Zheng and Dietz, 1998a). In contrast, “osmotically leaky” epithelium in D. polymorpha
results in very elevated excretory fluid production (Dietz
et al., 1995), requiring extremely high rates of epithelial
ion uptake to replace ions lost in voluminous excretory fluids (Dietz and Byrne, 1997). Thus, the main
354
Robert F. McMahon and Arthur E. Bogan
osmoregulatory adaptations of C. fluminea and D.
polymorpha are quite different, the former being reduced epithelial permeability, and that of the latter, increased water excretion and active epithelial ion uptake.
Hormones regulate osmotic control in freshwater
bivalves. Cyclic AMP (cAMP) stimulates active Na
uptake by unionoideans, while prostglandins inhibit it.
In contrast, prostglandin inhibitors stimulate N uptake (Dietz et al., 1982; Graves and Dietz, 1982;
Saintsing and Dietz, 1983). Serotonin stimulates tissue
accumulation of cAMP, increasing active Na uptake.
Thus, an antogonistic relationship between serotonin
and prostaglandins modulates adenylate cyclase-catalyzed cAMP stimulation of active Na uptake. Not
surprisingly, high concentrations of serotonin occur in
unionoidean gill nerve tracts (Dietz, 1985; Dietz et al.,
1992). Gill concentrations of the neurotransmitters,
dopamine and norepinephrine, greatly declined in
unionoideans salt-depleted in extremely dilute mediums, while serotonin was regulated at near-normal levels, suggesting that serotonin regulates Na uptake for
ionic balance. The circadian rhythms of Na uptake in
freshwater clams (Graves and Dietz, 1980; McCorkleShiley, 1982) may be mediated by an antagonistic serotonin / prostaglandin hormonal system (Dietz, 1985).
When cerebropleural or visceral ganglia were ablated, individuals of the Indian unionoidean, Lamellidens corrianus, rapidly lost osmoregulatory capacity
which was restored by injection of ganglia extracts, indicating that ganglion neurosectory hormones are involved in water balance (Lomte and Jadhav, 1981b).
The affinity of the pedal ganglion of A. cygena for
monoamines controlling ion / water balance is temperature dependent indicative of a seasonal component to
osmoregulation (Hiripi et al., 1982).
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
North American freshwater bivalves distributions,
particularly for unionoideans, have been well described. Species distribution maps for unionoideans
and sphaeriids exist for Canada (Clarke, 1973), the
United States (LaRocque, 1967a) and for the whole of
North American (Burch, 1975a, b; Parmalee and Bogan, 1998). LaRocque (1967b) describes living and
Pleistocene fossil assemblages at specific NA localities.
There is also a massive literature, too numerous to cite
here, describing species occurrences or species assemblages in various NA drainage systems.
Native (non-introduced) NA sphaeriid species have
broad distributions, often extending from the Atlantic
to Pacific coasts. Introduced to NA from southeast Asia
in early 1900s (McMahon, 1999), Corbicula fluminea
(i.e., light-colored shell morph of Corbicula) has a similarily widespread NA distribution, inhabiting drainages
on the west coast of the United States, the southern tier
of states, and throughout states east of the Mississippi
River, with the exception of the most northern states,
and into northern Mexico (McMahon, 1999) (Fig. 11).
A second, unidentified species of Corbicula (i.e., the
dark-colored shell morph) is restricted to isolated,
spring-fed drainages in southcentral Texas and southern California and Arizona (Fig. 11) (Britton and Morton, 1986). In contrast, NA unionoidean species generally have more restricted distributions. Few species
range on both sides of the continental divide and a
large number are limited to single drainage systems
(Burch, 1975b; LaRocque, 1967a).
1. Dispersal
Widespread NA distributions of sphaeriids and
Corbicula relative to unionoideans reflect fundamental
differences in their dispersal capacities. Unionoideans
depend primarily on host fish glochidial transport for
dispersal (Kat, 1984), their ranges reflecting those of
their host fish species (Haag and Warren, 1998). While
host fish glochidial transport increases probability of
dispersal into favorable habitats, as host fish and adult
unionoidean habitat preferences generally coincide
(Kat, 1984), it limits the extent of dispersal, leading to
highly endemic species. For example, electrophoretic
studies of peripheral Nova Scotian populations of
unionoideans suggest that they invade new habitats
mostly by host fish dispersal (Kat and Davis, 1984),
barriers to host fish dispersal being barriers to unionid
dispersal. Thus, distributions of modern and fossil NA
interior basin unionoidean assemblages are limited to
areas below major waterfall barriers to fish host upstream migration in the Lake Champlain drainage system of New York, Vermont, and Quebec (Smith,
1985a) and re-establishment of Anodonta implicata
populations in the upper Connecticut River drainage
closely followed restoration of its anadromous
glochidial clupeid fish host populations, by construction of fishways past numerous impoundments preventing upstream fish host dispersal (Smith, 1985b).
Vaughn and Taylor (2000) have found that 50% of
the variation in unionoidean assemblages is associated
with regional distribution and abundances of fishes, indicating that fish community structure is a determinant
of mussel community structure.
Sphaeriids and C. fluminea have evolved dispersal
mechanisms that make them more invasive than
unionoideans, accounting for their more cosmopolitan
distributions. Juvenile sphaeriids disperse between
11. Mollusca: Bivalvia
355
FIGURE 11
Distribution of Corbicula in the United States. Hatched area is the distribution of the
light-colored morph of Corbicula, Corbicula fluminea. The solid areas are the distribution of the
dark-colored morph of Corbicula, yet to be assigned a species designation.
drainage systems by clamping shell valves onto limbs of
aquatic insects, feathers of water fowl (Burky, 1983),
or even limbs of salamanders (Davis and Gilhen, 1982)
first noted by Darwin (1882). Some sphaeriids survive
ingestion and regurgitation by ducks, which commonly
feed on them, allowing long-distance dispersal (Burky,
1983). The rapid spread of C. fluminea through NA
drainage systems (McMahon, 1999), while partly anthropomorphically mediated, also resulted from its natural dispersal capacities. The long juvenile mucilaginous byssal thread or filamentous algae on which
juveniles settle becomes entangled in the feet or feathers of shore birds or water fowl, making them transport vectors (McMahon, 1999). Its natural dispersal
capacity has allowed it to spread into areas of northern
Mexico where human-mediated transport is highly unlikely (Hillis and Mayden, 1985), and into southern
Britain during interglacial periods (McMahon, 1999).
Dreissena polymorpha, byssally attaches to floating
wood or boat and barge hulls, facilitating long distance
transport (Mackie and Schloesser, 1996), and to
macrophytic vegetation that can be transported across
drainages by nesting shore birds and water fowl.
Juveniles of C. fluminea can be passively transported long distances downstream suspended in water
currents (McMahon, 1999). Water currents also disperse the planktonic veliger of D. polymorpha
(Mackie and Schloesser, 1996). Adult C. fluminea can
be carried downstream over substratum by water cur-
rents (Williams and McMahon, 1986), a process facilitated by production of a mucus dragline from the
exhalant siphon (Prezant and Charlermwat, 1984).
Passive hydraulic dispersal of juvenile and adult C. fluminea not only accounts for its extraordinary ability
to invade downstream portions of drainages after introduction (1999), but also leads to its impingement
and fouling of industrial, agricultural, and municipal
raw-water systems (1999). Similarly, current-mediated
transport of free-swimming veliger and passively suspended juvenile D. polymorpha, and of adults attached to floating substrates or carried as clumps of individuals over the bottom accounts for its dispersal in
European drainage systems after escape from the
Caspian Sea (Mackie and Schloesser, 1996). This
species rapidly spread downstream throughout the
Great Lakes and St. Lawrence River from its original
upstream introduction into Lake St. Clair in
1985 – 1986 (Mackie and Schloesser, 1996). It has also
been carried throughout most of the Mississippi River
and adjoining tributaries both by downstream hydrological transport and upstream by attachment to the
hulls of commercial barges (Mackie and Schloesser,
1996) (Fig. 12). Zebra mussels invaded the lower
Great Lakes, St. Lawrence River and Erie-Barge Canal
by 1990, later invading portions of the upper Great
Lakes, the Hudson River, and the Finger Lakes. D.
polymorpha entered the Mississippi Drainage from
Lake Michigan through the Illinois River and spread
356
Robert F. McMahon and Arthur E. Bogan
D. polymorpha, D. bugensis and the majority of
unionoideans are gonochoristic, requiring simultaneous
introduction of males and females to found a new population, thus reducing their capacity as invaders.
2. Anthropomorphic Impacts
FIGURE 12 Distribution of the zebra mussel, Dreissena polymorpha, in North America as of Spring, 2000.
downstream to the Mississippi Delta and upstream to
La Crosse, Wisconsin. It had invaded all major tributaries of the Mississippi River by 1998 with the exception of the Missouri River, where the first specimens
were reported upstream of its confluence with the
Platte River in 1999. As of early 2000, D. polymorpha
occupied drainages in 21 eastern and midwestern U.S.
states and in the Canadian provinces of Ontario and
Quebec, including 62 confirmed populations in small,
isolated inland lakes (Fig. 12). Dreissena bugensis, the
second dreissenid species occupying NA inland waters
is sympatric with D. polymorpha in Lakes Huron,
Erie, and Ontario and the western end of the ErieBarge Canal (New York Sea Grant, 1999).
Capacity for downstream transport and byssal attachment has made D. polymorpha a major NA biofouling pest species, recapitulating its history in Europe
(Claudi and Mackie, 1993). Juvenile sphaerids are also
passively, hydrologically transported downstream
(McKillop and Harrison, 1982) which may be an important dispersal mode for many species in this family.
In contrast, hydrological transport is extremely rare in
unionoideans (Imlay, 1982).
As both sphaeriids and C. fluminea are selffertilizing hermaphrodites (see Section II.B.5), a single
individual can found a new population. In contrast,
The diversity of native NA unionoid bivalves is
represented by two of the six recognized families of the
Unionoidea: Margaritiferidae with two genera and five
species; and Unionidae with 49 genera, 278 species, 13
subspecies (Turgeon et al., 1998; Johnson, 1998;
Williams and Fradkin, 1999). North American
unionoidean diversity represents 51 of the approximately 165 unionoidean genera, and between onefourth and one-third of the world’s unionoidean species
diversity (Bogan, 1993; Bogan and Woodward, 1992).
Neves et al. (1997) documented that 261 of these taxa
or 91% of NA unionoidean diversity occurs in the
southeastern United States where it is focused in the
Mobile Bay, Tennessee and Cumberland River basins.
Alabama, with parts of the Mobile Bay and Tennessee
River basins, has the greatest unionoidean diversity
with 175 taxa followed by Tennessee with 129 taxa.
This great diversity of unionoidean bivalves began
to be impacted as it began to be described, with European expansion across North America. It was noticed
quite early on that the unionoidean fauna was declining (Higgins, 1858). By the turn of the 20th century,
people were observing that the unionoidean fauna of
entire regions was being decimated or had disappeared.
Rhoads (1899) described extirpation of mussels from
the Monongahela River at Pittsburgh due to damming
and pollution. Ortmann (1909) noted the loss of the
freshwater mussels, crayfish and fish fauna from the
upper Ohio River Basin in western Pennsylvania due to
acid run-off from coal mines and the complete destruction of the Pigeon River unionoidean fauna in east Tennessee by pollution (Ortmann, 1918). Van der Schalie
(1938) warned that the Tennessee Valley Authority’s
construction of dams on the Tennessee River could lead
to long-term negative impacts on, and the eventual destruction of, its freshwater fauna.
Extinction of NA freshwater bivalve species was
not reported until Stansbery (1970, 1971) listed 11 presumed extinct taxa. Turgeon et al., (1988) listed 13
taxa, Williams et al., (1993) listed 12% of the unionoid
taxa as extinct, and most recently Turgeon et al., (1998)
listed 35 taxa as presumed extinct (Table II). As of
1997, 77 NA unionid taxa were endangered, 43 threatened and 73 of special concern with only 70 species
listed as currently stable and additional species being
added to federal and state lists yearly (Williams et al.,
1993; Neves et al., 1997). Massive historical losses of
unionoideans have been revealed by comparison of
11. Mollusca: Bivalvia
TABLE II
357
Extinct Freshwater Unionoidean Bivalves
Species
Common name
States of occurrence
Alasmidonta mccordi Athearn, 1964
Alasmidonta robusta Clarke, 1981
Alasmidonta wrightiana (Walker, 1901)
Elliptio nigella (Lea, 1852)
Epioblasma arcaeformis (Lea, 1831)
Epioblasma biemarginata (Lea, 1857)
Epioblasma flexuosa (Rafinesque, 1820)
Epioblasma florentina florentina (Lea, 1857)
Epioblasma haysiana (Lea, 1834)
Epioblasma lenior (Lea, 1842)
Epioblasma lewisii (Walker, 1910)
Epioblasma obliquata obliquata (Rafinesque, 1820)
Epioblasma personata (Say, 1829)
Epioblasma propinqua (Lea, 1857)
Epioblasma sampsonii (Lea, 1861)
Epioblasma stewardsonii (Lea, 1852)
Epioblasma torulosa gubernaculum (Reeve, 1865)
Epioblasma torulosa torulosa (Rafinesque, 1820)
Epioblasma turgidula (Lea, 1858)
Lampsilis binominata Simpson, 1900
Medionidus mcglameriae van der Schalie, 1939
Pleurobema altum (Conrad, 1854)
Pleurobema avellanum Simpson, 1900
Pleurobema bournianum (Lea, 1840)
Pleurobema chattanoogaense (Lea, 1858)
Pleurobema flavidulum (Lea, 1861)
Pleurobema hagleri (Frierson, 1900)
Pleurobema hanleyianum (Lea, 1852)
Pleurobema johannis (Lea, 1859)
Pleurobema murrayense (Lea, 1868)
Pleurobema nucleopsis (Conrad, 1849)
Pleurobema rubellum (Conrad, 1834)
Pleurobema troschelianum (Lea, 1852)
Pleurobema verum (Lea, 1861)
Quadrula tuberosa (Lea, 1840)
Coosa elktoe
Carolina elktoe
Ochlockonee arcmussel
Winged spike
Sugarspoon
Angled riffleshell
Leafshell
Yellow blossom
Acornshell
Narrow catspaw
Forkshell
Catspaw
Round combshell
Tennessee riffleshell
Wabash riffleshell
Cumberland leafshell
Green blossom
Tubercled blossom
Turgid blossom
Lined pocketbook
Tombigbee moccasinshell
Highnut
Hazel pigtoe
Scioto pigtoe
Painted clubshell
Yellow pigtoe
Brown pigtoe
Georgia pigtoe
Alabama pigtoe
Coosa pigtoe
Longnut
Warrior pigtoe
Alabama clubshell
True pigtoe
Rough rockshell
AL
NC, SC
FL
AL, GA
AL, KY, TN
AL, KY, TN
AL, IL, IN, KY, OH,TN
AL, KY, TN
AL, KY, TN, VA
AL, TN
AL, KY, TN
AL, IL, IN, KY, OH, TN
IL, IN, KY, OH
AL, IL, IN, KY, OH,TN
IL, IN, KY
AL, KY, TN
TN, VA
AL, IL, IN, KY, OH,TN, WV
AL, AR, TN
AL, GA
AL
AL, GA
AL
OH
AL, GA, TN
AL
AL
AL, GA, TN
AL
AL, GA, TN
AL, GA
AL, GA, TN
AL, GA, TN
AL
TN, VA
(From Williams et al., 1998b.)
present species assemblages with those of earlier surveys
or with recent fossil assemblages (Neves and Zale,
1982; Parmalee and Klippel, 1982, 1984; Parmalee
et al., 1982; Ahlstedt, 1983; Havlik, 1983; Stern, 1983;
Hoeh and Trdan, 1984; Miller et al., 1984; Hartfield
and Rummel, 1985; Taylor, 1985; Mackie and Topping,
1988; Nalepa and Gauvin, 1988; Starnes and Bogan,
1988; Baily and Green, 1989; Bogan, 1990; Anderson
et al., 1991; Counts et al., 1991; Hornbach et al., 1992;
Parmalee and Hughes, 1993; Hoke, 1994; Blalock and
Sickel, 1996). Extirpations of sphaeriid faunae are far
less common, but have occurred (Paloumpnis and Starrett, 1960; Mills et al., 1966). Unionoidean assemblages
in Indian middens near the upper Ohio River yielded at
least 32 species, while a 1921 survey yielded only 25 of
the midden species, and a 1979 survey, only 13 of the
midden species in the same area, indicative of massive
species extirpation (Taylor and Spurlock, 1982). Similar
historical loss of Indian midden species has occurred in
the Tennessee River Drainage (Parmalee, 1988). Further
evidence of environmental change in the upper Ohio
River includes recent establishment of 15 unionid
species previously unreported on Indian middens or earlier surveys (Taylor and Spurlock, 1982).
Such historical data clearly indicate the toll that
anthropomorphic activities are taking on the NA
unionoidean fauna (Neves, 1993; Neves et al., 1997).
The United States Fish and Wildlife Service began listing unionoideans as threatened and endangered after
passage of the Endangered Species Act of 1973. By
1998, 56 taxa were listed as endangered (Turgeon
et al., 1998), with about 70% of NA unionoideans at
some level of imperilment (Williams et al., 1993).
Bogan (1998) reviewed the causes for decline of NA
freshwater bivalve diversity and attributed it, and
species extinctions, to habitat destruction (loss of both
358
Robert F. McMahon and Arthur E. Bogan
unionoidean and host fish habitat), pollution including
acid mine runoff, pesticides, heavy metals, commercial
exploitation and introduced species. Bogan (1997)
summarized this:
“The central cause of this decline and decimation of the
freshwater molluscan fauna is the modification and destruction
of their aquatic habitat, with sedimentation as a leading major
factor. Sources of sedimentation include poor agricultural and
timbering practices. Damming of major rivers has also had a
dramatic impact on this fauna with the loss of unionid obligate
host fish due to changes in local water quality and loss of
habitat. In-stream gravel mining, dredging, and canalization
have further eliminated stable aquatic habitat. Acidic mine
drainage and various point and non-point pollution sources also
continue to decimate local aquatic mollusk populations.”
Evidence of negative anthropomorphic impacts on
unionoidean populations is extensive. The freshwater
pearling industry can extirpate entire populations (Laycock, 1983), overfishing for pearls being a major factor
in the decline of the pearl mussel, Margaritifera magaritifera in Great Britain (Young and Williams, 1983a).
Commercial unionid shell fisheries that provide seed
pearls for the marine cultured pearl industry negatively
impact NA populations (Williams et al., 1993). Impoundments of rivers slow flow and allow accumulation of silt, leading to mussel fauna reductions (Duncan
and Thiel, 1983; Parmalee and Klippel, 1984; Stern,
1983; Starnes and Bogan, 1988; Blay, 1990; Williams
et al., 1992; Houp, 1993; Parmalee and Hughes,
1993). Impoundments also eliminate glochidial fish
hosts (Mathiak, 1979) or prevent fish host dispersal
of glochidia. Release of cold, hypolimnetic water
from impoundments negatively impacts downstream
unionoidean populations (Ahlstedt, 1983; Clarke,
1983; Vaughn and Taylor, 1999). Controlled water releases from impoundments lead to major flow-rate oscillations, either scouring the bottom of suitable mussel
substrates during high flows or causing lethal aerial exposure during low flows (Miller et al., 1984; Vaughn
and Taylor, 1999). Channelization of drainage systems
for navigation or flood control is detrimental to
unionoideans. Increased flow velocity and propeller
wash elevate suspended solids, which interfere with
mussel filter feeding and O2 consumption (Aldridge
et al., 1987; Payne and Miller, 1987). It reduces availability of stabilized sediments, sand bars, and low-flow
areas, all preferred unionoidean habitats (Payne and
Miller, 1989; Strayer and Ralley, 1993; Strayer, 1999a).
Pollution adversely affects bivalves. Unionoidean
faunas can be negatively impacted or extirpated by industrial pollution (Zeto et al., 1987; Wade et al.,
1993), urban wastewater effluents (sewage, silt, pesticides) and resultant eutrophication and hypoxia (Gunning and Suttkus, 1985; St. John, 1982; Neves and
Zale, 1982; Arter, 1989; Strayer, 1993), silt and acid
discharges from mines (Taylor, 1985; Warren et al.,
1984; Anderson et al., 1991) siltation from bank erosion due to deforestation, destruction of riparian zones,
and poor agricultural practice (Hartfield, 1993;
Williams et al., 1993) or disturbance and silt from
river-bed gravel mining (Brown and Curole, 1997). Advent of modern sewage treatment on the Pearl River,
Louisiana, allowed re-establishment of five previously
absent unionid species (Gunning and Suttkus, 1985).
Nonindigenous species (i.e., “biological pollution”)
present a new threat to NA unionoidean taxa. The
nonindigenous zebra mussel, D. polymorpha, byssally
attaches in great numbers to the exposed, posterior,
siphonal shell regions of unionoideans, leading to their
slow starvation as the zebra mussels strip suspended
food particles from their inhalant current. Thus, zebra
mussel infestations have extirpated a number of
unionoidean species populations from the lower Great
Lakes (Schloesser et al., 1996).
3. Physical Factors
Physical factors influence bivalve distributions.
While environmental requirements are species specific,
a number of generalities appear warranted. Sediment
type clearly affects distribution patterns. Unionoideans
are generally most successful in areas where flow is
moderate with a stable substrate of coarse sand or
sand – gravel mixtures, and are generally absent from
substrates with heavy silt loads and very low water
flow (Strayer and Ralley, 1993; Strayer, 1999a). In the
Wisconsin and St. Croix rivers, only 7 of 28 unionid
species occurred in sand – mud sediments, the majority
preferring sand – gravel mixtures. Only three species,
Pyganodon grandis, Lampsilis teres, and L. siliquoidea,
preferred sand – mud substrates (Stern, 1983). Some
unionoidean species select preferred sediment types
(Baily, 1989). In rivers subjected to periodic high flows,
unionoideans oriented with siphons facing upstream to
a greater extent than those in stable flow rivers, a position which presents the narrowest profile to the current, reducing chances of flow-induced dislodgment
(Di Maio and Corkum, 1997). The relatively specific
substrate and flow requirements of many unionoidean
species may account for their patchy and clumped distributions (Strayer, 1999a). Courser sediments allowing
free exchange of interstitial and surface waters may be
a requirement for early survival of recently excysted juvenile unionoideans with low pH, hypoxia, and ammonia accumulation in interstitial water being correlated
with juvenile mortality in sediments where free water
exchange is reduced (Buddensiek et al., 1993). Thus,
sediment limitations for juvenile development may
result in patchy or clumped adult unionoidean distributions. In contrast, C. fluminea is able to colonize
11. Mollusca: Bivalvia
359
FIGURE 13
Sediment relationships in sphaeriid clam communities from sites along a depth transect in
Britannia Bay, Ottawa River, Canada. (A) Mean sphaeriid density (Shannon – Weaver d) values for various
sphaeriid communities in relation to mean sediment geometric particle size. (B) Mean sphaeriid density values
at various depths. Vertical bars about points are 95 percent confidence limits. Note increase in diversity with
decrease in mean sediment particle size and maximization of density at depths less than one meter. (Redrawn
from data of Kilgour and Mackie, 1988.)
habitats ranging from bare rock through gravel / sand
to silt (McMahon, 1999), which has allowed it to invade a wide variety of NA drainage systems. Its optimal habitat is oxygenated sand or gravel – sand (Belanger et al., 1985).
In contrast to unionoideans, species diversity in
Pisidium increases with decreasing particle size (Fig.
13A), becoming maximal at a mean particle diameter
of 0.18 mm (Kilgour and Mackie, 1988). In southeastern Lake Michigan, Pisidium density and diversity were
maximal in very fine sand – clay and silt – clay sediments, while peak Sphaerium diversity occurred at
somewhat larger particle sizes (Zbeda and White,
1985). Thus, there are differences in substrate preferences among sphaeriids, perhaps associated with sediment organic detritus feeding mechanisms in Pisidium
(see Section III.C.2).
Apparent differences in substrate preferences may
be associated with species-specific differences in optimal water velocities. Unionoideans are most successful
where velocities are low enough to allow sediment
stability, but high enough to prevent excessive siltation
(Strayer and Ralley, 1993; Strayer, 1999a), making
well-oxygenated, coarse sand and sand – gravel beds
optimal habitats for riverine species with such habitats
being more critical for survival of juvenile
unionoideans than adults (Buddensiek et al., 1993).
Low or variable velocities allow silt accumulations that
make sediments too soft for maintenance of position in
adult unionoideans (Lewis and Riebel, 1984; Salmon
and Green, 1983) or which interfere with their filter
feeding and gas exchange (Aldridge et al., 1987). In
contrast, periodic scouring of substrates during high
flows removes substrate and unionoideans, and prevents their successful resettlement (Young and
Williams, 1983b). Indeed, unionoidean densities decline in areas of high flow (Way et al., 1990b). Sediment type did not affect burrowing in three lotic
unionid species (P. grandis, Elliptio complanata, and
Lampsilis siliquoidea) (Lewis and Riebel, 1984), suggesting that it is not involved in substrate preferences,
but unionoideans may move to preferred sediments
(Baily, 1989). C. fluminea, with its relatively heavy,
ridged shell and rapid burrowing ability, is better
adapted for life in high current velocities and unstable
substrates than most unionoideans (McMahon, 1999).
In the Tangipahoa River, Mississippi, it colonizes unstable substrates from which unionoideans are excluded (Miller et al., 1986). In contrast to the majority
of unionoideans and C. fluminea, many sphaeriid
species occur in small ponds and the profundal portions of large lakes, where water flow is negligible and
the substrate has high silt contents and heavy organic
loads (Burky, 1983). Preference of some sphaeriids for
low-flow, silty habitats may reflect interstitial sediment
detritus feeding, particularly in Pisidium (Lopez and
Holopainen, 1987) (see Section III.C.2). Byssal attachment allows D. polymorpha to inhabit relatively high
flow areas compared to other bivalves (pediveliger
larvae settle in flows up to 1.5m / sec). It also makes it
successful epibenthic species in lentic habitats with a
preponderance of hard substrates from which native
NA bivalves are generally eliminated (Claudi and
Mackie, 1993).
360
Robert F. McMahon and Arthur E. Bogan
Water depth affects freshwater bivalve distributions.
Most unionoideans prefer shallow habitats less than
4 – 10 m deep (Stone et al., 1982; Salmon and Green,
1983; Machena and Kautsky, 1988; Way et al., 1990b),
although some species occur in deeper lotic waters if
well oxygenated. C. fluminea is also restricted to shallow, near-shore lentic habitats (McMahon, 1999), as are
the majority of Sphaerium and Musculium species (Fig.
13B) (Zbeda and White, 1985; Kilgour and Mackie,
1988). In contrast, some species of Pisidium inhabit
profundal regions of lakes (Holopainen and Jonasson,
1983; Kilgour and Mackie, 1988). Depth distributions
of D. polymorpha vary between habitats; however,
adults rarely occur above 2 m and dense populations extend to 4 – 60 m, but are restricted to well-oxygenated
waters above the epilimnion. The quagga mussel, D.
bugensis, extends to greater, oxygenated depths than
D. polymorpha in the Great Lakes (Mills et al., 1996),
but is unlikely to penetrate hypoxic hypolimnetic waters
as it is less hypoxia tolerant than D. polymorpha (P. D.
Johnson and R. F. McMahon, unpublished data). Recently settled juveniles of D. polymorpha migrate to
deeper water (Mackie and Schloesser, 1996), perhaps
avoiding wave-induced agitation (Clarke and McMahon, 1996b) or ice scour.
Limitation of most lentic bivalves to shallow habitats may reflect their poor hypoxia tolerance. In lentic
habitats, hypolimnetic waters are often hypoxic. As
many species of unionoideans, Sphaerium and Musculium (Burky, 1983), Dreissena (McMahon, 1996)
and C. fluminea (McMahon, 1999) cannot maintain
normal O2 uptake under severely hypoxic conditions,
they are mostly restricted to shallow, well-oxygenated
habitats. Juvenile unionids are less hypoxia-tolerant
than adults (Dimock and Wright, 1993), preventing
colonization of hypoxic habitats. In contrast, many
species of Pisidium are extreme V̇O2 regulators ( Burky,
1983) allowing them to inhabit highly hypoxic
hypolimnetic habitats (Jonasson, 1984a, b). However,
summer hypoxia retards growth and reproduction in
profundal Pisidium populations, indicating that hypoxia can deleteriously impact even hypoxia-tolerant
species (Halopainen and Jonasson, 1983). Hypoxiaintolerant C. fluminea invaded the profundal regions of
a small lake only after artificial aeration of its hypoxic
hypolimnetic waters (McMahon, 1999). Sewageinduced hypoxia in the Pearl River, Louisiana, extirpated its unionoidean fauna (Gunning and Suttkus,
1985). Even hypoxia-tolerant profundal Pisidium communities are extirpated by extreme hypoxia (Jonasson,
1984a).
Ambient pH does not greatly limit the distribution
of freshwater bivalves. The majority of species prefer
waters of pH above 7.0; species diversity declines in
acidic habitats (Okland and Kuiper, 1982). However,
unionoideans can grow and reproduce over a pH range
of 5.6 – 8.3, a pH of less than 4.7 – 5.0 being the absolute lower limit (Fuller, 1974; Okland and Kuiper,
1982; Kat, 1982; Hornbach and Childers, 1987). Low
pH may have sublethal effects on unionoideans. It
reduces shell thickness (Hinch et al., 1989), tissue cholesterol content (Rao et al., 1987), and hemolymph
concentrations of Na, K, and Cl (Pynnönen, 1991;
Mäkelä and Oikari, 1992). Unionoidean glochida and
juveniles are less low pH tolerant than adults (Huebner
and Pynnönen, 1992; Dimock and Wright, 1993), perhaps preventing colonization of mildly acidic waters. In
contrast, Pyganodon grandis showed no change in hemolymph Na, K, and Cl concentrations after transfer into an acidic lake (pH 5.9) from an alkaline lake
(Malley et al., 1988). Some sphaeriids are relatively insensitive to pH or alkalinity. Species richness, growth
and reproduction in sphaeriid faunas of six lowalkalinity lakes was similar to those in higher alkalinity
lakes (Rooke and Mackie, 1984a, b; Servos et al.,
1985). Musculium partumeium and Pisidium casertanum inhabited acid lakes in New York State (pH
6.0) (Jokinen, 1991). Indeed, maximal laboratory
growth and reproduction in Musculium partumeium
occurred at pH 5.0, suggesting adaptation to moderately acidic habitats (Hornbach and Childers, 1987).
Low-pH habitats generally have low calcium concentrations. Low pH leads to shell dissolution and
eventual mortality if shell penetration occurs (Kat,
1982). Sphaeriids occur in waters with Ca concentrations as low as 2 mg Ca / L, while the unionid, Elliptio
complanata, occurs at 2.5 mg Ca / L (Rooke and
Mackie, 1984a). Among freshwater bivalves, D. polymorpha is the most calciphilous and pH intolerant;
adults requiring pH 6.5 and 12 mg Ca / L and
veliger larvae requiring pH 7.4 and 24 mg Ca / L
for successful development (McMahon, 1996). Freshwater bivalves actively take up Ca2 at medium concentrations as low as 0.5 mM Ca / L (0.02 mg Ca / L,
see Section II.A.1), which is far below their minimal
ambient Ca2 concentration of 2 – 2.5 mg / L. Thus, the
minimum ambient calcium concentration appears to be
that at which the rate of calcium uptake and deposition
to the shell exceeds that of calcium loss from shell dissolution and diffusion, allowing maintenance of shell
integrity and growth. As many factors affect shell deposition and dissolution rates (e.g., temperature, pH,
and calcium concentration), the minimal Ca concentration and / or pH tolerated by a species may vary between habitats dependent on interacting biotic and
abiotic parameters and are often species-specific. Low
calcium waters usually have low concentrations of
other biologically important ions, making them inhos-
11. Mollusca: Bivalvia
pitable to bivalves even if Ca concentrations are suitable for shell growth.
Temperature influences bivalve distributions;
species have specific upper and lower limits for survival and reproduction (Burky, 1983). For example,
intolerance of 2°C prevents C. fluminea from colonizing drainages in the northcentral United States,
which reach 0°C in winter (Fig. 11) (McMahon,
1999), leading to low-temperature winter-population
kills on the northern edge of its range (Sickel, 1986).
Thus, many northern U.S. C. fluminea populations are
restricted to areas receiving heated effluents (McMahon, 1999). In contrast, the maximal temperature for
development of D. polymorpha eggs and larva is 24°C
and adults do not tolerate temperatures 30°C, preventing this species from colonizing southern and
southwestern U.S. waters which exceed 30°C in summer (McMahon, 1996). Indeed, in the most southern
U.S. states, D. polymorpha only occurs in the lower
Mississippi River which rarely exceeds 30°C (Hernandez et al., 1995).
Water-level variation affects bivalve distributions.
Declining water levels during droughts or dry periods
expose relatively immotile bivalves for weeks or
months to air. Restriction of many bivalve populations
to shallow waters makes them susceptible to emersion.
Many sphaeriid species and some unionoidean taxa are
highly tolerant of air exposure, surviving prolonged
seasonal emersion in ephemeral or variable level habitats (Burky, 1983; White, 1979; Byrne and McMahon,
1994). These species display unique emersion adaptations (Byrne and McMahon, 1994 and Section II.C.4).
Freshwater bivalve distribution is also related to
stream size or order. Unionoidean species diversity increased with distance downstream in the Sydenhan
River, Ontario (Mackie and Topping, 1988). Similarly,
stream size proved to be the only useful predictor of
unionoidean species richness of six tested factors
(i.e., stream size, stream gradient, hydrologic variablity, Ca concentration, physiographic province and
presence / absence of tides) in the Susquehanna,
Delaware and Hudson River drainages (Strayer, 1993).
The number of unionoidean species in Michigan
drainage systems increased proportionately with system size; mainly by species additions (Strayer, 1983)
(Fig. 14), but species richness could not be completely
accounted for by drainage area size (note the high degree of variation in species richness versus drainage
area values in Fig. 14), suggesting that other environmental variables affect unionoidean distribution patterns (Strayer, 1993). In the Ohio River drainage, the
number of fish species was directly related to drainage
area, and the number of unionoidean species directly
related to the number fish species, suggesting that
361
FIGURE 14
Unionoidean mussels species richness (total number of
species present at a particular site) as a function of stream size measured by its total drainage area in km2 in southeastern Michigan,
United States. Numbers next to points indicate the number of observations falling on that point. The relationship between drainage area
and mussel species richness was statistically significant r 0.68,
P 0.001). (Redrawn from Strayer, 1983.)
unionoidean diversity is partially dependent on fish
glochidial host species availability (Watters, 1993).
Similarly, Margaritifera hembeli populations with the
highest juvenile recruitment rates occurred in stream
reaches with the highest densities of its fish host (Johnson and Brown, 1998). Among other variables affecting species richness are stream hydrology, affected by
surface geology and soil porosity. Porous soils retain
water, buffering runoff, so that streams draining them
have constant flows and rarely dry, allowing them to
support greater mussel diversity than streams draining
soils of poor water-infiltration capacity that are prone
to flooding – drying cycles. In variable-flow streams,
emersion or low oxygen in stagnant pools during dry
periods, and bottom scouring and high silt loads during floods reduce species richness (Strayer, 1983). Indeed, flow stability, high O2, reduced flood risk, and
low silt loads of larger streams appear to account for
their increased bivalve species richness (Fig. 14)
(Strayer, 1983, 1993; Way et al., 1990b; Strayer and
Ralley, 1993). However, some unionoidean species,
such as Amblema plicata, Pyganodon grandis and Fusconaia flava, are adapted to small, variable flow
streams (Strayer, 1983; Di Maio and Corkum, 1995)
while other species favor tidally influenced portions of
rivers (Strayer, 1993).
The above physical factors may interact to determine distributions of bivalves. The density of the
unionoidean, M. hembeli, was affected by stream order,
hardness, depth, substrate size and compaction, and
362
Robert F. McMahon and Arthur E. Bogan
flow rate, with highest densities occurring in shallow
areas of second-order streams, of hardness 8 mg / L,
with higher flow rates, and stable sediments with larger
particle sizes (Johnson and Brown, 2000).
B. Reproduction and Life History
North American freshwater bivalves display extraordinary variation in life history and reproductive
adaptations. Life-history traits (e.g., those affecting reproduction and survival, including growth, fecundity,
life span, age to maturity, and population energetics)
have been reviewed for freshwater molluscs (Calow,
1983; Russell-Hunter and Buckley, 1983) and specifically for freshwater bivalves (Burky, 1983; Mackie,
1984; Mackie and Schloesser, 1996; McMahon, 1999).
Most research involves the Sphaeriidae, with less information for unionoideans. Sphaeriids are good subjects
for life-history studies, because of their greater abundances, ease of collection and laboratory maintenance,
relatively simple hermaphroditic life cycles, ovovivipar-
ity, semelparity, release of completely formed miniature
adults, and relatively short life spans. In contrast,
unionoideans are more difficult subjects because they
are gonochoristic, long-lived, interoparous, often rare
and difficult to collect, and have life cycles complicated
by the parasitic glochidial stage. The life-history traits
of C. fluminea and D. polymorpha have been intensely
studied due to their invasive nature and economic importance as fouling organisms (Mackie and Schloesser,
1996; Nichols, 1996; McMahon, 1999).
1. Unionoidea
The life-history characteristics of freshwater
unionoideans are clearly different from those of
sphaeriids, C. fluminea or D. polymorpha (Table III).
The majority of unionids live in large, stable aquatic
habitats which buffer them from periodic catastrophic
population reductions, typical of smaller, unstable
aquatic environments (see Section III.A). In stable
habitats, long-lived adults accumulate in large numbers
(Payne and Miller, 1989). A few species inhabit ponds
TABLE III Summary of the Life History Characteristics of North American Freshwater Bivalves, Unionoidea, Sphaeriidae,
Corbicula fluminea, Driessena polymorpha
Life history traita
Unionoidea
Sphaeriidae
Corbicula fluminea
Dreissena polymorpha
Life span (years)
6 – 100 (Species
dependent)
6 – 12
1 – 5 (species dependent)
1–4
4–7
0.17 – 1.0 (1 year
in some species)
Hermaphroditic
025 – 0.75
0.5 – 2
Hermaphrodictic
self-fertilizing
Rapid throughout
life
35,000
Gonochoristic
Age at maturity (years)
Reproductive mode
Fecundity (young per avg.
adult per breeding season)
Gonochoristic (few
hermaphroditic species)
Rapid prior to maturity,
slower thereafter
200,000 – 17,000,000
per female
Juvenile size at release
Very small, 50 – 450 m
Slow relative to Unionoideans,
C. fluminea or D. polymorpha
3 – 24 (Sphaerium)
2 – 136 (Musculium)
3 – 7 (Pisidium)
Large 600 – 4150 m
Relative juvenile survivorship
Relative adult survivorship
Extremely low
High
High
Intermediate
Semelparous or iteroparous
Highly iteroparous
Number of reproductive
efforts per year
Assimilated energy
respired (%)
Nonrespired energy
allocated to growth (%)
Nonrespired energy allocated
to reproduction (%)
Turnover time in days ( mean
standing crop biomass
: biomass per day ratio)
Habitat stability
One
Growth rate
a
Very small 250 m
Rapid throughout
life
30,000 – 40,000
per female
—
Semelparous or Iteroparous
(species dependent)
1 – 3 (continuous
in some species)
21 – 91 (Avg. 45%)
Extremely low
Low 2 – 41%
per year
Moderately
iteroparous
Two (spring
and fall)
11 – 42
Extremely small,
40 m
Extremely low
Intermediate 26 – 88%
per year
Moderately
iteroparous
One (2 – 8 months
long)
—
85.2 – 97.5
65 – 96 (Avg. 81%)
58 – 71
96.1
2.8 – 14.8
4 – 35 (Avg. 19%)
15
4.9
1790 – 2849
27 – 1972 (generally 80)
73 – 91
53 – 869 (habitat
dependent)
Stable
Intermediately stable
Unstable
Moderately unstable
See text for literature citations to data on which this table was based.
11. Mollusca: Bivalvia
363
FIGURE 15 Anatomic features of freshwater bivalve larval stages. (A) the D-shaped juvenile of Corbicula
fluminea, the freshwater Asian clam (shell length 200 m). (B) The glochidium larva of unionoideans, which
is parasitic on fish. Depicted is the glochidium of Pyganodon characterized by the presence of paired spined
hooks projecting medially from the ventral edges of the shell valves and an attachment thread; not all
unionoidean species have glochidia with these characteristics (50– 400 m in diameter depending on species).
(C) Lateral and anterior views of the free-swimming, planktonic veliger stage of Dreissena polymorpha, the
zebra mussel; the veliger is 40 –290 m in diameter and uses the ciliated velum to swim and feed on phyto- and
bacterioplankton. Presence of a foot as shown in this diagram indicates development to the settlement competent pediveliger stage. The juveniles of freshwater sphaeriid species are large and highly developed, having
essentially adult features (Fig. 2A) at birth.
(Burch, 1975b), but their life-history traits have not
been studied.
An important aspect of the unionoidean life-history
traits is their parasitic glochidium larval stage (Fig.
15B). With the exception of Simpsonaias ambigua,
whose glochidial host is the aquatic salamander, Necturus maculosus, all other known NA unionoideans
have glochidial fish hosts (Watters, 1994b). The significance of glochidia unionoidean reproduction has been
reviewed by Kat (1984) and details of glochidial incubation and development in marsupial brood pouches
were described in Section II.B.5. The glochidium has a
bivalved shell adducted by a single muscle. Its mantle
edge contains sensory hairs. In the genera Unio, Anodonta, Pygauodon, Megalonaias, and Quadrula, a
long threadlike structure projects from the mantle beyond the ventral valve margin which may be involved
with detection of and / or attachment to, fish hosts.
There are three general forms of glochidia. In the
subfamily Anodontinae, “hooked glochidia” occur, with
triangular valves from whose ventral edges project an inward curving hinged hook covered with smaller spines
(Fig. 15B). On valve closure, the hooks penetrate the
skin, scales, or fins of fish hosts, allowing glochidial attachment and encystment on host external surfaces. The
majority of NA unionoideans produce “hookless
glochidia” with more rounded valves bearing reinforcing
structures and / or small spines or stylets on their ventral
margins which generally attach and encyst on fish gills.
“Axe-head glochidia” of the genus Potamilus have a
flared ventral valve margin, and near-rectangular valves
which may have hooklike structures on each corner.
Their host attachment sites are unknown (Kat, 1984).
Glochidia initially attach to fish by clamping (snapping) the valves onto fins, scales, and / or gill filaments.
They are not host-specific in attachment, attaching to
any contacted fish (Kat, 1984). In contrast, gravid
adult unionids (European Anodonta anatina) appear
to release glochidia in response to chemicals released
by host fish (Jokela and Palokamgas, 1993). Once
364
Robert F. McMahon and Arthur E. Bogan
released, glochidia display ‘snapping’ behavior’ (i.e.,
host attachment behavior), the valves being rapidly and
repeatedly adducted. In M. margaritifera, glochidial
valve snapping is stimulated by the presence of mucus,
blood, gill tissue, or fins of their brown trout host, but
not by water currents or tactile stimulation (Young and
Williams, 1984a), suggesting use of chemical cues to
detect and attach to host fish.
Glochidia encyst in fish host tissues within 2 – 36 h
of attachment and may or may not grow during encystment, depending on species. Time to juvenile excystment is also species-dependent, ranging from 6 to 160
days, but is reduced at higher temperatures (Zale and
Neves, 1982; Kat, 1984). Unsuitable host fish reject
glochidia after encystment (Kat, 1984), fish blood
serum components dictating host suitability (Neves et
al., 1985). Since glochidia nonselectively attach to fish
(Kat, 1984; Neves et al., 1985), host suitability appears
more dependent on fish immunity than on glochidial
host recognition. Indeed, even suitable host fish reject
glochidia, rejection rate being host fish species-dependent (Haag et al., 1999). Numbers of M. margaritifera
glochidia encysted in a brown trout population declined with time (Young and Williams, 1984b), with
laboratory studies revealing that only 5 – 12% of M.
margaritifera glochidia successfully excyst from infected host fish, indicating host rejection of most encysted individuals (Young and Williams, 1984a).
Unionoideans have adaptations that increase likelihood of glochidia contact with fish hosts. Glochidial release occurs annually, but duration of release is speciesdependent with some species having more than one
annual release period (Fukuhara and Nagata, 1988;
Gordon and Smith, 1990). Cycles of gametogenesis and
glochidial release may be controlled by neurosecretory
hormones (Nagabhushanam and Lomte, 1981). Tachytictic mussels are short-term breeders, whose glochidial
development and release occurs between April and August; shedding of glochidia often corresponding with either migratory periods of anadromous fish hosts, or
nesting periods of their host fish. Fish hosts of tachytictic unionoideans often construct nests in areas harboring dense unionoidean populations, where fish host nest
construction by fanning away substrates, and fish host
fanning of developing embryos provide optimal conditions for glochidial – host contact. Thus, a high proportion of nest-building fish species, such as centrarchids,
are glochidial hosts for NA unionoideans (Watters,
1994b). In contrast, bradytictic unionoidean species are
long-term breeders, retaining developing glochidia in
gill marsupia throughout the year, releasing them in
summer (Kat, 1984).
When released from adult mussels, glochidia are
generally bound by mucus into discrete packets, which
either dissolve, releasing glochidia, or remain intact as
discrete “conglutinates” of various species-specific
forms and colors. Glochidia with attachment threads
(Fig. 15B) are released in tangled mucus threads that
dissolve relatively rapidly. Many of these glochidia
possess hooks and attach to fish-host external surfaces.
Hooked glochidia are larger than other types of
glochidia (Bauer, 1994). In some unionoideans, mucilaginous networks of glochidia persist, enhancing host
contact by suspending glochidia above the substrate.
Unionids with hookless glochidia that attach to fish
gills may release conglutinates that mimic the food
items of their fish hosts. They resemble brightly colored
oligochaetes, flatworms, or leeches. Some species hold
their wormlike conglutinates partially extruded from
the exhalant siphon, making them more obvious to fish
hosts. Some Lamsilis species, such as, Lampsilis perovalis, produce a “supercongultinate” that contains all
the glochidia in the marsupial gill and resembles a small
fish. It is tethered to the female’s exhalant siphon by a
long, transparent mucus strand (Fig. 16A). In flowing
water, it mimics the darting motions of a small fish, eliciting attacks by host fish (Haag et al., 1995). Consumption of such conglutinates releases glochidia within the
buccal cavity of the fish followed by transport to gill
filament attachment sites on ventilatory currents.
The most unusual unionoidean host food mimicry
involves pigmented muscular, posterior mantle edge extensions in female Lampsilis and Villosa. These “mantle flaps’ (Fig. 16B) resemble the small fish or macroinvertebrate prey of their fish hosts (Haag and Warren,
1999). Gravid females extend the posterior shell margins well above the substrate and pulsate the projecting
mantle flaps to mimic a small, actively swimming fish
or moving macroinvertebrate. When fish strike these
mantle lures, glochidia are forcibly released through
posterior pores in the marsupial gill (often projected
between the mantle flaps, Fig. 16B) assuring glochidial
ingestion and contact with the fish host gills (Kat,
1984; Haag and Warren, 1999).
As the glochidia of some unionoideans do not grow
while encysted, their parasitic nature has been questioned. However, in vitro glochidial culture experiments
suggest that they absorb organic molecules from fish tissues and require fish plasma for development and metamorphosis (Isom and Hudson, 1982), indicative of a
true host – parasite relationship. Indeed, glochidial infection damages fish hosts, especially juvenile fish (Cunjak
and McGladdery, 1991; Panha, 1993).
Like other parasites, glochidia are shed in huge
numbers to ensure the maximum host contact and attachment. Unionoidean fecundity ranges from 10,000
to 17,000,000 glochidia per female per breeding season
(Parker et al., 1984; Young and Williams, 1984b;
365
FIGURE 16
Reproductive adaptations in female Lamsilid unionoideans. (A) Superconglutinate tethered on a mucus strand extending from the exhalant siphon of Lampsilis
perovalis. Glochidia are encapsulated in the flattened terminal lure (detailed in the upper
portion of the figure) at the end of the mucus strand which makes darting fishlike movements in water currents, enticing host fish to strike the lure, thus assuring release of
glochidia into its buccal cavity for attachment to the fishes’ gills. (Redrawn from Haag et
al., 1995) (B) The modified, mantle flaps of a female specimen of Lampsilis ovata, which
mimic the small fish prey (note eye spot and lateral linelike pigmentation) of its predatory host fish species. The posterior portions of the marsupial outer demibranchs are
projected from the mantle cavity to lie between the flap lures. When the mantle flap lures
are struck by an attacking host fish, glochidia are released through pores in the marsupial gill ensuring their entry into the Figh’s buccal cavity and attachment to its gills. Superconglutinates and mantle flap lures are characteristic of the unionoidean genus,
Lampsilis.
366
Robert F. McMahon and Arthur E. Bogan
Paterson, 1985; Paterson and Cameron, 1985; Jansen
and Hanson, 1991; Bauer, 1994), brood size increasing
exponentially with female size (Janson and Hanson,
1991; Bauer, 1994). Probability for glochidial survival
to metamorphosis is extremely small. In a natural
population of M. margaritifera, a species that does not
produce glochidial conglutinates, only 0.0004% of released glochidia successfully encysted in fish hosts. Of
these, only 5% were not rejected before excystment;
and, of those successfully metamorphosing, only 5% established as juveniles in the substrate (Young and
Williams, 1984b). Thus, only one in every 100,000,000
shed glochidia became settled juveniles. High glochidial
mortality makes the effective fecundity of unionoideans
extremely low, which is not unusual for a species
adapted to stable habitats. Similarly, only 0.007% of released glochida of Pyganodon grandis encysted in their
host fish (Perca flavescens), however, once encysted,
they had relatively high survival, 27% surviving to, and
excysting after, two years (Jansen and Hanson, 1991).
Based on these data and the fecundity ranges listed in
Table III, only 0.002 – 0.17 of the glochidia produced by
a female unionoidean would successfully settle as juveniles in sediment. Therefore, the main advantages of the
glochidial stage appear to be a directed dispersal by fish
hosts into favorable habitats (Kat, 1984) and utilization
of fish-host energy resources to complete development
to a juvenile large enough to effectively compete after
settlement in adult habitats (Bauer, 1994).
There is a trade-off between glochidial size and
number of glochidia released. Species with large
glochidia (including toothed glochidia) produce fewer
glochidia while species with small glochidia release
them in great numbers (Bauer, 1994). Species releasing
fewer, larger glochidia, tend to have a greater range of
host-fish species and prefer lower flow habitats (Bauer,
1994). Production of many small glochidia occurs in
species with limited fish hosts and may increase chances
of glochidial – host contact, while having a wide range
of host-fish species allows production of fewer, larger
glochidia, because chances of glochidium contact with a
suitable host are higher. In the highly K-selected, stable
habitats of most unionoideans, excysting at a large size
makes juvenile mussels more competitive, increasing
likelihood of survival to maturity. Thus, small glochidia
of unionoideans, such as Margaritifera, remain encysted
for longer periods (1000 days), allowing growth to a
more competitive size before excystment. In contrast,
among species with larger glochidia, the encystment period is much shorter (200 – 300 days) before attainment
of competitive juvenile size. The long encystment periods of small glochidia reduce their chances for successful excystment due to increased chances of host mortality and / or rejection by the host’s immune system,
selecting for production of greater numbers of glochidia
to assure recruitment of enough juveniles to sustain
population size (Bauer, 1994).
Glochidial utilization of fish-host energy stores prevents their direct competition with adults for limited
food and space resources (Bauer, 1994), as occurs in juvenile C. fluminea and D. polymorpha. Glochidial parasitism also allows female unionoideans to devote relatively small amounts of nonrespired, assimilated energy
to reproduction (2.8 – 14.5% of total nonrespired, assimilated energy), leaving the majority for somatic
tissue growth (Table V) (Negus, 1966; James, 1985;
Paterson, 1985). Allocation of a high proportion of
energy to tissue growth is characteristic of species
adapted to stable habitats (i.e., K-selected species). As
unionoideans are often very long-lived (Margaritifera
margaritifera up to 130 – 200 years) and highly
iteroparous (6 – 10 reproductive periods throughout
life), allocation of the majority of nonrespired energy
to growth increases probability of adult survival to
future reproductive efforts. Increased growth rate and
reduction of reproductive effort increases competitiveness and lowers probability of predation and / or mortality associated with reproductive effort or dislodgment from the substrate during floods. All these
characteristics increase unionoidean fitness in stable
habitats (Sibly and Calow, 1986; Bauer, 1994).
In the majority of unionoideans, greatest shell
growth occurs in immature individuals during the first
4 – 6 years of life (Hanson et al., 1988; Payne and
Miller, 1989; Harmon and Joy, 1990) (Fig. 17A). Indeed, relative shell growth in young unionids is greater
than in sphaeriids or C. fluminea and D. polymorpha
(Table III). Unionoidean shell-growth rate declines exponentially with age, but tissue biomass accumulation
rate remains constant or increases with age (Fig. 17A,
B; see also Haukioja and Hakala, 1978). Thus, early in
life, increases in shell size and biomass occur preferentially over tissue accumulation; whereas after maturity
(6 years), shell growth slows and tissue accumulates
at proportionately higher rates. Delayed maturity in
unionoideans (6 – 12 years, Table III) allows allocation
of all nonrespired assimilation to growth early in life
and, due to an exponential relationship between size
and fecundity, leads to reproductive effort being primarily sustained by the oldest, largest individuals
(Downing et al., 1993).
Growth rates in unionoideans are affected by abiotic conditions. Shell growth of field-enclosed specimens P. grandis declined with depth, being greatest
3 m (Hanson et al., 1988). However, in natural populations, depth did not influence shell growth in P.
grandis (Hanson et al., 1988) or A. woodiana (Kiss
and Pekli, 1988), suggesting that vertical migration
11. Mollusca: Bivalvia
367
FIGURE 17 The shell and tissue growth of selected species of unionids
(Anodonta anatina, open circles; Unio pictorum, open squares; Unio tumidus,
open tiangles; Elliptio complanata, solid circles; and Pyganodon cataracta, solid
squares). (A) Mean shell-length increase with increasing age over the entire life
span of each species. (B) Mean dry tissue-weight increase with increasing age over
the entire life span of each species. Note that increase in shell length declines with
age, while dry tissue weight increases either linearly or exponentially with age.
(Data from Negus, 1966; Paterson, 1985; and Paterson, and Cameron, 1985.)
may mask depth influences on growth in free-living individuals (Hanson et al., 1988). Unionoidean growth
rates also vary seasonally with level of precipitation
(Timm and Mutvei, 1993) and can be depressed by
pollutants (Harman and Joy, 1990).
Once mature, large adult unionoideans display high
age-specific survivorship between annual reproductive
efforts, being 81 – 86% in 5 – 7 year-old Anodonta
anatina (Negus, 1966), greater than 80% in mature M.
margaritifera (12 – 90 years) (Bauer, 1983), with a lower
value of 15.6% in Pleurobema collina (Hove and
Neves, 1994). High adult survivorship, long life spans,
and low juvenile survivorship account for the preponderance of large adult individuals in natural
unionoidean populations (Bauer, 1983; James, 1985;
Negus, 1966; Paterson, 1985; Paterson and Cameron,
1985; Tevesz et al., 1985; Hansen et al., 1988; Huebner
et al., 1990; Woody and Holland-Bartels, 1993; Hove
and Neves, 1994; Vaughn and Pyron, 1995). Populations dominated by adults are characteristic of stable,
highly competitive habitats (Sibly and Calow, 1986).
The preponderance of large, long-lived adults in
unionoidean populations causes them to have high proportions of standing crop biomass relative to biomass
production. The relationship between standing crop
biomass and biomass production rate can be expressed
as turnover time, computed in days as the average
standing crop of a population divided by its mean daily
productivity rate (Russell-Hunter and Buckley, 1983).
Long-lived unionids, with populations dominated by
large adults, have extremely long turnover times
ranging from 1790 – 2849 days (computed from data in
James, 1985; Negus, 1966; Paterson, 1985; Huebner
et al., 1990) compared with sphaeriids (27 – 1972 days),
C. fluminea (73 – 91 days), or D. polymorpha (53 –
869 days) (Table III). Such extended turnover times
368
Robert F. McMahon and Arthur E. Bogan
characterize long-lived, iteroparous species from stable
habitats (Russell-Hunter and Buckley, 1983).
In only one aspect do unionoideans deviate from the
life-history traits expected of species inhabiting stable
habitats and experiencing extensive competition (i.e.,
K-selected). That is in the production of large numbers
of small young (glochidia). However, as described
above, this is an adaptation ensuring sufficiently high
probability of glochidial – host fish contact. Thus, species
producing conglutinates resembling fish host prey items
have greater glochidial – host contact and produce fewer
(200,000 – 400,000 glochidia per female) and larger
glochidia (Kat, 1984). In contrast, M. margaritifera,
which releases very small, dispersed glochidia, has extraordinarily high fecundities (up to 17,000,000 glochidia
per female) (Young and Williams, 1984b).
Extended life spans, delayed maturity, low effective
fecundities, reduced dispersal, high habitat selectivity,
poor juvenile survival and extraordinarily long
turnover times make unionoidean populations highly
susceptible to human perturbations. Because of their
life-history traits (particularly long life spans and low
effective fecundities), they do not recover rapidly once
decimated by pollution or other human- or naturallymediated habitat disturbances (see Section III.A). Successful juvenile settlement appears particularly affected
by disturbance, with population structures indicating
periods when entire annual generations are not recruited (Bauer, 1983; Negus, 1966; Payne and Miller,
1989). Reduction in population densities may reduce
fertilization success in unionoideans. Complete failure
of fertilization in Elliptio complanata occurred at densities 10 mussels / m2 with densities 40 mussels / m2
required for 100% of females to be fertilized (Downing
et al., 1993). Thus, anthropomorphically influenced reduction of unionoidean population densities to low levels could prevent further recruitment. Such disturbance-induced lack of juvenile recruitment raises the
specter of many NA unionoidean populations being
composed of dwindling numbers of long-lived adults
destined for extirpation as anthropomorphic disturbances prevent reproduction and / or juvenile recruitment in aging adult populations (for an example, see
Vaughn and Pyron, 1995).
2. Sphaeriidae
The Sphaeriidae display great intra- and interspecific
life history variation (Holopainen and Hanski, 1986;
Mackie, 1984; Way, 1988). Like unionoideans, their lifehistory traits do not fall into suits associated with stable
or unstable habitats. Instead, they include the short life
spans, early maturity, small adult size, and increased
energetic allocation to reproduction associated with
adaptation to unstable habitats, and the slow growth
rates, low fecundity, and release of large, fully developed
young associated with adaptation to stable habitats
(Sibly and Calow, 1986) (Table III). Sphaeriids are euryoecic (i.e., have a broad habitat range); some members
inhabiting stressful habitats, such as ephemeral ponds,
and small, variable flow streams, while others live in stable, profundal lake habitats (Burky, 1983). Here, we
generalize the life-history traits of sphaeriids within
adaptive and evolutionary frameworks. However, the
degree of inter- and intraspecific life-history variation in
this group is such that, for each generality, specific exceptions can be cited (Mackie, 1979).
Central to understanding sphaeriid life-history
traits is their viviparous reproduction (Mackie, 1978).
All species brood embryos in specialized chambers
formed from evaginations of the exhalant side of the
inner demibranch gill filaments. Maternal nutrient material is supplied to embryos supporting their growth
and development to release as fully formed miniature
adults (see Section II.B.5). Thus, even though adult
sphaeriids are the smallest of all NA freshwater bivalves, they release the largest young (Mackie, 1984).
Among 13 sphaeriid species, average birth shell length
ranged from 0.6 to 4.15 mm (Burky, 1983; Holopainen
and Hanski, 1986; Hornbach and Childers, 1986;
Hornbach et al., 1982; Mackie and Flippance, 1983a),
much larger than unionoidean glochidia (0.05 – 0.45
mm, Bauer, 1994), C. fluminea juveniles (0.25 mm,
McMahon, 1999), or D. polymorpha veliger larvae
(0.04 – 0.07 mm, Nichols, 1996) (Table III). Based on
shell lengths, newborn sphaeriids have (3.4 – 2.1) 105
times greater biomass than newborns of other groups.
Ratios of maximum adult shell length : birth shell
length in sphaeriids range from 2.8:1 to 5.4:1, suggesting that newborn juveniles have biomass 0.6 – 4.6% of
the maximum biomass of adults. There is a significant
direct relationship between these two parameters,
shown in Figure 18.
Large offspring size reduces sphaeriid fecundity.
Average clutch sizes range from 3 to 24 per young per
adult for Sphaerium, 2 to 136 per young per adult for
Musculium, and 1.3 to 16 per young per adult for
Pisidium (Burky, 1983; Holopainen and Hanski,
1986). Even with reduced fecundity, large juvenile biomass requires allocation of large amounts of nonrespired energy to reproduction (x 19%, Burky,
1983) compared to unionoideans (14.8%), C. fluminea (15%) or D. polymorpha (4.9%) (Table III). As
developing juveniles have high metabolic rates (Burky,
1983; Hornbach, 1985; Hornbach et al., 1982) and are
supported by adult energy stores (Mackie, 1984), estimates of reproductive costs based on released juvenile
biomass may grossly underestimate actual reproductive
costs in this group.
11. Mollusca: Bivalvia
FIGURE 18 The relationship between shell length (SL) of juveniles
at birth and maximal SL of adults for 13 species of sphaeriid freshwater clams. Note that juvenile birth length increases linearly with
maximal adult size, suggesting that adult size may limit juvenile birth
size in sphaeriids. The solid line represents the best fit of a linear
regression relating birth size to maximal adult size as follows: Birth
SL 0.0387 0.236 mm (maximal adult SL in mm) (n 13, r
0.935, F 76.3, P 0.0001). (Data from Holopainen and Hanski,
1986; Hornbach and Childers, 1986; Hornbach et al., 1982; and
Mackie and Flippance, 1983a.)
Life-history hypotheses predict that the low fecundity and large birth size of sphaeriids would maximize
fitness in stable habitats. However, the majority of
sphaeriids inhabit variable ponds and streams or profundal habitats subject to hypoxia (Burky, 1983;
Holopainen and Hanski, 1986). Burky (1983) and Way
(1988) have argued that the harsh conditions of such
habitats are seasonally predictable, making them, in reality, stable. Thus, sphaeriid species inhabiting them
have adaptations that prevent catastrophic population
reductions during seasonal episodes of environmental
stress (see Section II.C). Thus, sphaeriid populations approach carrying capacity, leading to selection for lifehistory adaptations associated with the intense intraspecific competition characteristic of life in stable habitats.
Seasonal water-level fluctuation and hypoxia often
more severely impact smaller individuals. Thus,
production of large, well-developed juveniles by
sphaeriids may increase their probability of surviving
predictable environmental stress. As such stress can
cause high adult mortality (Burky et al., 1985b;
Holopainen and Jonasson, 1983; Hornbach et al.,
1982; Jonasson 1984b), fitness would also be increased
369
by devoting greater proportions of nonrespired assimilation to any one reproductive effort, as chances of
adult survival to the next reproduction are low. Thus,
sphaeriids devote relatively higher proportions of
energy to reproduction than other freshwater bivalves
(Burky, 1983) (Table III). This combination of r- selected and K-selected traits, optimizing sphaeriid fitness
in habitats with periodic, predictable stress, is a lifehistory strategy called “bet hedging” (Stearns, 1980).
The early maturation, characteristic of many
sphaeriid species (Burky, 1983; Holopainen and
Hanski, 1986) (Table III), is due to the advanced development of newborn juveniles, and allows reproduction
before onset of seasonal environmental stress. This trait
is particulary displayed by sphaeriid species inhabiting
ephemeral ponds and streams. They have rapid growth,
early maturity, and reduced numbers of reproductive efforts. Thus, Musculium lacustre, Pisidium clarkeanum,
and P. annandalei from temporary drainage furrows in
Hong Kong live less than 1 year, and have only one or
two life-time reproductive efforts (Morton, 1985,
1986). Similarly, Musculium lacustre, Pisidium casertanum, P. variable, Spharium fabule, and Musculium
securis had only 1 – 3 reproductive efforts within life
spans of 1 to 2 years (Mackie, 1979) and a lake species,
S. japonicum, is univoltine and semelparous (Park and
Okino, 1994). An ephemeral pond population of Musculium partumeium lived 1 year, was semelparous, reproduced just prior to pond-drying, and devoted a large
proportion of nonrespired assimilation (18%) to reproduction (Burky et al., 1985b). Ephemeral pond populations of M. lacustre were similarly univoltine and semelparous, devoting 19% of nonrespired energy to
reproduction (Burky, 1983). A Sphaerium striatinum
population, subject to flooding, lived 1 year or less and
reproduced biannually (Hornbach et al., 1982), devoting 16.1% of nonrespired assimilation to reproduction
(Hornbach et al., 1984b). In contrast, when populations of these species occur in more permanent habitats,
they become bivoltine, reproducing in both spring and
fall. Spring generations are iteroparous, reproducing in
the fall and following spring; whereas fall generations
are semelparous, reproducing only in the spring
(Mackie, 1979; Burky, 1983; Burky et al., 1985b).
Some Pisidium species live in the profundal portions of large, more permanent lentic habitats where
they tolerate periodic hypoxia after summer formation
of the hypolimnion (see Section II.C.3). As these
profundal species are tolerant of hypoxia, such environments are more stable than small ponds and
streams, although far less productive, reducing food
availability (Holopainen and Hanski, 1986). Thus,
profundal sphaeriids display life-history strategies
differing from those in small ponds and streams
370
Robert F. McMahon and Arthur E. Bogan
(Burky, 1983; Holopainen and Hanski, 1986). Profundal populations have reduced growth rates, longer life
spans of 4 to 5 years, delayed maturation often exceeding 1 year, higher levels of iteroparity, and univoltine
reproductive patterns (Holopainen and Hanski, 1986),
all life-history traits characteristic of more stable habitats (Sibly and Calow, 1986). Interestingly, shallowwater and profundal populations of the same species of
Pisidium may display different life-history tactics.
Compared to profundal populations, shallow-water
populations grow more rapidly, mature earlier, have
shorter life spans, and tend toward semelparity
(Holopainen and Hanski, 1986), life-history traits associated with unstable habitats. As pisidiids have welldeveloped dispersal capacities, this variation in lifehistory tactics among species’ populations occupying
different habitats in unlikely to result from genetic
adaptation. Rather, much of this variation appears to
be environmentally induced plasticity and is reflected in
the highly variable turnover times reported for sphaeriids (range 27 – 1972 days, Table III).
Freshwater bivalve growth rates are highly dependent on ecosystem productivity. Populations from productive habitats have greater assimilation, allowing allocation of greater absolute amounts of nonassimilated
energy to growth (Burky, 1983). Shallow, freshwater
habitats are usually highly productive, warm, and
rarely O2-limited and, thus, support higher growth
rates. Conversely, profundal environments are less productive, cooler, and often O2-limited, leading to lower
growth rates. Since sphaeriids mature at a species-specific size irrespective of growth rate (Burky, 1983;
Holopainen and Hanski, 1986), rapid growth leads to
early maturity in shallow-water habitats and slow
growth to delayed maturity in profundal habitats.
Sphaeriids also die at species-specific terminal sizes
whether terminal size is attained rapidly or slowly. As
growth rate determines the time required to reach terminal size, fast-growing, shallow-water individuals
reach terminal sizes more rapidly (often within less
than 1 year) allowing participation in only one or two
reproductive efforts, while slow-growing, profundal individuals (Holopainen and Hanski, 1986) reach terminal size more slowly, allowing participation in a greater
number of reproductive efforts. Similar interpopulation
growth rate and reproductive variation has been
recorded in several NA sphaeriid species (Mackie,
1979). The fundamentally ecophenotypic nature this
variation has been demonstrated for Pisidium casertanum. When individuals were reciprocally transferred
between two populations with different life-history
traits or co-reared in the laboratory, the majority of
life-history trait differences proved to be environmentally induced. However, electrophoresis revealed ge-
netic differences between populations and transfer and
laboratory corearing demonstrated a small portion of
observed variation to be genetically based (Hornbach
and Cox, 1987). Extensive ecophenotypic plasticity
may account for the euryoecic nature and cosmopolitan distributions of sphaeriids (see Section III.A). Certainly, capacity to adjust growth rates, maturity, reproductive cycles, life cycles, and energetic allocation
patterns to compensate for habitat biotic and abiotic
variation enables sphaeriids to have relatively wide
niches and, thus broad distributions.
3. Corbicula fluminea
The introduced Asian freshwater clam, Corbicula
fluminea, unlike unionoideans and sphaeriids, has lifehistory traits adapting it to unstable, unpredictable
habitats (McMahon, 1999), making it the most invasive
of all NA freshwater bivalves. C. fluminea grows
rapidly, in part because it has higher filtration and assimilation rates than other bivalves (Foe and Knight,
1986a; Lauritsen, 1986a; Way et al., 1990a). In a natural population, a relatively small proportion of assimilation (29%) was devoted to respiration (Table V), the
majority (71%) being allocated to growth and reproduction (Aldridge and McMahon, 1978). Laboratory
studies have also showed that 59 – 78% (Lauritsen,
1986a) or 58 – 89% of assimilation (Foe and Knight,
1986a) was allocated to tissue production. Thus, C. fluminea has among the highest net production efficiencies
recorded for any freshwater bivalve, reflected by its low
turnover times, ranging from 73 to 91 days (Table III).
The very high proportion of nonrespired assimilation (85 – 95%) devoted to growth in C. fluminea
(Aldridge and McMahon, 1978; Long and McMahon,
unpublished data), sustains rapid shell growth (shell
length 15 – 30 mm in the first year of life, 35 – 50
mm in the terminal third-to-fourth year) (McMahon,
1983a). High growth rates decrease probability of
predation, as fish and bird predators feed only on
small individuals (Section III.C.3) thus, increasing
probability of survival to next reproduction in this
moderately iteroparous species. Increase in shell size
occurs at the expense of tissue production during summer maximal growth (Long and McMahon, unpublished data), suggesting that larger shells optimize fitness. High growth rates sustain the highest population
production rates (10.4 – 14.6 g organic carbon / m2 per
year, Aldridge and McMahon, 1978) reported for any
freshwater bivalve (Burky, 1983), with dense populations producing 1000 – 4500 g C / m2 per year
(McMahon, 1999).
Newly released juveniles are small (shell length
⬇250 m), but completely formed, with a characteristically D-shaped shell, adductor muscles, foot, statocysts,
11. Mollusca: Bivalvia
gills, and digestive system (Kraemer and Galloway,
1986) (Fig. 15A). Juveniles are denser than water and
settle and anchor to sediments or hard surfaces with a
single mucilaginous byssal thread. However, they are
small enough to be suspended in turbulent water currents and hydrologically transported (McMahon,
1999). A relatively low amount of nonrespired assimilation is allocated to reproduction (5 – 15%; Aldridge and
McMahon, 1978; Long and McMahon, unpublished
data), equivalent to that of unionids, but less than
sphaeriids (19%, Table III). However, the species’ elevated assimilation rates allow higher actual allocation
to reproduction than in other species.
As the juvenile of C. fluminea is small (organic carbon biomass 0.136 g), fecundity is large, ranging
from 97 to 570 juveniles per adult per day during reproductive efforts, yielding an average annual fecundity
of 68,678 juveniles per adult per year (McMahon,
1999). Early juvenile survivorship is extremely low and
mortality rates remain high throughout life (74 – 98%
in the first, 59 – 69% in the second, and 93 – 97% in the
third year) (McMahon, 1999), causing populations to
be dominated by juveniles and immatures. High adult
mortality and population dominance by immature individuals characterizes species adapted to unstable habitats (Stearns, 1980).
The majority of NA C. fluminea populations have
two annual reproductive periods (i.e., are bivoltine)
(McMahon, 1999). It is hermaphroditic and capable of
self-fertilization (Kraemer and Galloway, 1986; Kraemer et al., 1986) such that single individuals can found
new populations. Spermiogenesis occurs only during
reproductive periods, but gonads contain mature eggs
throughout the year (Kraemer and Galloway, 1986).
C. fluminea matures within 3 – 6 months at a shell
length of only 6 – 10 mm (Kraemer and Galloway,
1986). Thus, juveniles born in spring can grow to maturity and participate in reproduction the following fall
(McMahon, 1999) (Table III). Maximum life span is
variable between populations and within populations,
ranging from 1 to 4 years (McMahon and Williams,
1986a; McMahon, 1999). Early maturity allows this
iteroparous species to participate in 2 – 7 reproductive
efforts, depending on life span.
A relatively short life span, early maturity, high fecundity, bivoltine reproduction, high growth rates,
small juvenile size, and capacity for downstream dispersal make C. fluminea both highly invasive and
adapted for life in truly unstable, disturbed lotic habitats subject to unpredictable catastrophic faunal reductions. Its high reproductive potential and growth rates
allow it to rapidly attain high densities after invading a
new habitat or after catastrophic population declines.
Thus, it is successful in NA drainage systems subject to
371
periodic anthropomorphic interference such as channelization, navigational dredging, sand and gravel
dredging, commercial and / or recreational boating, and
organic and / or chemical pollution, compared to less
resilient unionoideans or sphaeriids (McMahon, 1999).
Surprisingly, C. fluminea is more susceptible to environmental stresses, such as temperature extremes,
hypoxia, emersion, and low pH than most sphaeriids
and unionoideans (McMahon, 1999), making it highly
susceptible to human disturbance. With only a limited
capacity to tolerate unpredictable environmental stress,
why is C. fluminea so successful in disturbed habitats?
The answer lies in its ability to recover from disturbance-induced catastrophic population crashes much
more rapidly than either sphaeriids or unionids. C. fluminea rapidly re-establishes populations even if disturbance has reduced them to a few widely separated individuals, as all individuals are hermaphrodites capable of
self-fertilization and have high fecundities. Downstream
dispersal of juveniles from viable upstream populations
allows rapid re-establishment of decimated populations,
the accelerated growth, high fecundity, and relatively
short life spans of this species allowing recovery to normal age – size distributions and densities within 2 – 4
years (McMahon, 1999). Biannual (bivoltine) reproduction also increases its probability of surviving catastrophic density reductions, as it prevents loss of an entire generation to chance environmental disturbance
(“bet hedging,” Stearns, 1980). Capacity for rapid recolonization allows C. fluminea to sustain populations
in substrates subject to periodic flood scouring from
which slower growing and late maturing unionoideans
are eliminated (Way et al., 1990b, Section III.B.1).
Like sphaeriids, NA C. fluminea populations display high interpopulation variation in life-history traits
(McMahon, 1999). As there is little or no genetic variation among NA populations (McLeod 1986), this variation must be ecophenotypic. Growth rates increase
and time to maturity and life spans decrease in populations from more productive habitats (McMahon,
1999). On the northern border of its NA range, low
temperatures reduce growth and reproductive periods,
such that populations are univoltine rather than bivoltine. A slow growing C. fluminea population was univoltine and semelparous in an oligotrophic Texas lake
(McMahon, unpublished data). Even within populations, life-history tactics vary, dependent on year-toyear variations in temperature and primary productivity (McMahon and Williams, 1986a; Williams and
McMahon, 1986). As in sphaeriids, ecophenotypic lifehistory trait variation is adaptive in C. fluminea, allowing colonization of a variety of habitats. Thus, this
species is highly euryoecic and highly invasive, making
it the single most successful and economically
372
Robert F. McMahon and Arthur E. Bogan
costly aquatic animal introduced to North America
(McMahon, 1999).
4. Dreissena Polymorpha
The zebra mussel, Dreissena polymorpha, is the
most recently introduced NA bivalve species (Mackie
and Schloesser, 1996). Like C. fluminea, its life-history
characteristics (McMahon, 1996; Nichols, 1996) make
it highly invasive. Unlike other NA bivalves, it releases
sperm and eggs making fertilization completely external. Developments leads to a free-swimming planktonic
veliger larva (Fig. 15C) which feeds and grows in the
plankton for 8 – 10 days before settlement (Nichols,
1996). The planktonic veliger enhances the zebra mussel dispersal ability. Veligers in the Illinois River traveled at least 306 km downstream before settlement
(Stoeckel et al., 1997). Numbers of dispersing veligers
can be astounding. Stoeckel et al. (1997) estimated that
veligers in the Illinois River reached densitiess 250
veligers / L which resulted in veligers passing a fixed
point at a rate as 75 106 per second. Total annual
veliger flux was estimated to be 1.935 1014 and
2.131 1014 per year in 1994 and 1995, respectively.
Adults byssally attached to floating objects are also
transported downstream.
Zebra mussels sexually mature in the first or second year of life (first year in most NA populations)
have life spans of 3 – 5 years in Europe and 2 – 3 years
in North America (Mackie and Schloesser, 1996) and,
like C. fluminea, sustain high growth rates throughout
life (Fig. 19). It is iteroparous and univoltine; an individual participating in 3 – 4 annual reproductive periods within its life span. Reproduction is initiated above
10 – 12°C, but is maximized above 18°C. Spawning is
dependent on abiotic and biotic conditions, producing
peaks of veliger density within a reproductive season
(McMahon, 1996; Nichols, 1996; Ram et al., 1996;
Stoeckel, et. al. 1997). The freshly hatched veliger is
small (diameter 40 – 70 m), but the pediveliger
grows to 180 – 290 m just prior to settlement
(Nichols, 1996), leading to a 100 to 400-fold biomass
increase during the 8 – 10 day planktonic period.
Maximal D. polymorpha adult size ranges from
3.5 to 5 cm depending on growth rate, which, like terminal size, is dependent on habitat primary productivity and temperature. Growth rate of caged mussels in
Lake St. Clair, Michigan, initially enclosed at an average shell length (SL) of 4.2 m was 0.095 mm per day
over the first 150 days reaching a mean SL of 14.3 mm
and, thereafter, slowing down to a mean of 0.015 mm
per day for the next 182 days reaching a mean SL of
17.0 mm (Bitterman et al., 1994). Growth rates of
young mussels in the lower Mississippi River were temperature-dependent, peaking at 0.06 – 0.08 mm per day
FIGURE 19
Shell-growth rates in European populations of the
zebra mussel, Dreissena polymorpha. Growth rates for North American populations in Lake Erie are similar to or greater than faster
growing British populations depicted in this figure.
at ⬇16 – 17°C and declining at higher temperatures
(Allen et al., 1999), a result supporting laboratory
studies showing that increasing metabolic demands reduce assimilated energy allocation to growth above
15°C (Walz, 1978a).
Like C. fluminea, D. polymorpha allocates a very
high percentage (96.1%) of nonrespired assimilation to
somatic growth, leaving only 3.9% for reproduction
(Mackie et al., 1989; Table III); however, Sprung
(1991) reports a 30% postspawning reduction in female body mass, indicating higher reproductive energy
allocation in some populations. Allocation of a large
proportion of nonrespired assimilation to growth allows individuals to rapidly increase in size, making
them more competitive and less subject to predation.
Zebra mussel veligers settle on the shells of established
individuals, forming thick mats or clusters many shells
deep, in which competition for space and food must be
intense. Thus, rapid postsettlement juvenile growth is
highly adaptive as it allows stable byssal attachment to
the substrate and positioning of siphons at the mat surface, where food and O2 are most available. In spite of
the low levels of energy devoted to reproductive effort
by D. polymorpha, its very small eggs make individual
fecundity large, ranging from 30,000 to 106 eggs per
female (Sprung, 1991; Mackie and Schloesser, 1996).
Dreissena polymorpha population densities range
from 7000 to 114,000 individuals / m2 and standing
crop biomasses from 0.05 – 15 kg / m2 (Claudi and
11. Mollusca: Bivalvia
Mackie, 1993). These high densities and biomasses result from the tendency of juveniles to settle on substrates inhabited by adults forming dense mats or
clumps many individuals thick. High individual growth
rates and population densities lead to very high productivities, estimated to be 0.05 – 15 g C / m2 per year in
European populations, and ⬇75 g C / m2 per year in
NA Great Lakes populations based on dry tissue mass
productivity values in Mackie and Schloesser (1996).
The NA productivity value for D. polymorpha is still
relatively low compared to values of 1000 – 4500 g
C / m2 per year estimated for dense NA C. fluminea
populations (McMahon, 1999). Population growth
and productivity are highly habitat-dependent in D.
polymorpha, making turnover times variable, ranging
from low values of 53 days (highly productive) to high
values of 869 days (low productivity; Table III).
A high growth rate throughout life, elevated fecundity, short life span, and a capacity for adult and larval
down-stream dispersal make Dreissena (like Corbicula)
a highly invasive species. However, unlike C. fluminea,
D. polymorpha populations are restricted to more stable habitats such as larger, permanent lakes and rivers.
Its apparent preference for stable habitats is reflected
by its endemic range in the Caspian Sea and Ural River
(large stable habitats), avoidance of shallow, near-shore
lentic- and low-flow lotic habitats, relatively long age
to maturity (generally at least 1 year of life), iteroparity, gonochorism, and relatively high adult survivorship
(26 – 88% per year; Mackie and Schloesser, 1996).
Restriction of the original range of D. polymorpha
to the Caspian Sea and Ural River also suggests a limited dispersal capacity between drainage systems. Its
dispersal through Europe occurred only in the nineteenth century (and continues in western Asia today)
as interconnenting canal systems transported it between drainages. Dispersal of D. polymorpha between
drainages is primarily by human vectors, including
transport of adults attached to boat / barge hulls, ballast water dumping, and transport of veligers through
canal systems interconnecting catchments. Adults are
poorly tolerant of emersion (McMahon, 1996), precluding extensive natural overland dispersal unless human-mediated. Thus, dispersal of this species between
NA catchments has been primarily by human vectors,
with larger, permanent, navigable bodies of water being
most susceptible (Mackie and Schloesser, 1996). Without anthropomorphic vectors, dispersal of D. polymorpha through NA drainages would almost certainly have
proceeded at a much slower pace than recorded for C.
fluminea, but human activities have already lead to this
species being widely distributed in major NA drainages
east of the Mississippi River (Ram and McMahon,
1996; New York Sea Grant, 1999).
373
C. Ecological Interactions
1. Behavioral Ecology
Other than borrowing (see Section II.A.2), information on bivalve behavior is sparse. Reviews of molluscan neurobiology and behavior (Willows, 1985, 1986)
have no bivalve references. Lack of information, rather
than lack of complex and intriguing behaviors, reflects
difficulties in making behavioral observations on predominantly sessile bivalves surrounded by a shell.
There are a number of reproductive behaviors in
freshwater bivalves, including those ensuring unionioidean glochidial contact with fish hosts (Section II.B).
Adult C. fluminea display downstream dispersal behavior during reproductive periods. While juvenile clams
(SL 2 mm) are hydrologically transported throughout
the year, immatures (SL 2 – 7 mm) and adults (SL 7
mm) leave the substrate to be passively hydrologically
transported over the sediment surface (“rolling”) only
prior to reproductive periods (Fig. 20). Dispersing
adults have lower dry tissue weights, lower tissue organic carbon-to-nitrogen ratios (Williams and McMahon, 1989), higher levels of ammonia excretion, and
reduced molar O2 consumption-to-N2 excretion ratios
than those remaining in the substrate (Williams and
McMahon, 1985), all indicative of poor nutritional
condition. Thus, downstream dispersal allows starving
individuals to disperse from areas of low food availability and high intraspecific competition into areas
more nutritionally favorable for reproductive efforts
(Williams and McMahon, 1986, 1989).
Some adult unionoidean bivalves and C. fluminea
display surface locomotory behavior involving the
same movements of foot and valves described in Section II.B.2 for burrowing. Unionoidean surface locomotion is fairly common (Imlay, 1982), leaving tracts
in sediments 3 – 10-m long (Golightly, 1982). The adaptive significance of bivalve surface locomotion is not
well understood as it could attract potential predators.
However, it may be involved with pedal feeding on organic sediment deposits (see Section III.C.2).
Some species, such as Pyganodon grandis, migrate
vertically with seasonal changes in water level (White,
1979) to avoid emersion. Other species, such as
Uniomerus tetralasmus, C. fluminea, and some sphaeriids, remain in position and suffer prolonged emersion
(see Section II.C.4). In Texas, fire ants, Solenopsis
invicta, kill emersed bivalves (McMahon, unpublished),
suggesting that this introduced insect may be a new
threat to unionoideans and sphaeriids as it expands its
range in the southeastern U.S.
Some sphaeriids, C. fluminea, and D. polymorpha
also crawl on hard substrates or macrophytes.
Sphaerium corneum holds the valves erect and crawls
374
FIGURE 20 Seasonal variation in downstream dispersal behavior by juvenile, subadult and adult
Corbicula fluminea in the water intake canal of a power station. (A) Rate of impingement of adults dispersing downstream onto traveling screens in front of water intake embayments (shell length 10 mm).
(B) Rate of retention of subadults (shell length 1 – 7 mm) in a clam trap held on the substrate surface
of the intake canal. (C) Juveniles suspended in the water-column (shell length 2 mm). Right vertical
axis represents density of juveniles in the intake canal water column (solid circles connected by dashed
lines). Left vertical axis is number of juveniles entrained daily with intake water (open circles connected
by solid lines). Note that the adult and subadult clams display maximal downstream dispersal behavior
during only two periods, March – May and September – November). Juveniles occurred in the water column throughout the year; peak juvenile water column densities occurred during reproductive periods
and midwinter periods of low ambient water temperature. (From Williams and McMahon, 1986.)
11. Mollusca: Bivalvia
375
TABLE IV The Effects of Temperature on the Percentage of Time Spent in Various
Valve Movement Behaviors in Emersed Specimens of Corbiucla fluminea a
Temperature (°C)
Time with valves
closed (%)
Time with mantle
edge exposed (%)
Time with mantle edge
parted or attempting
to burrow (%)
15
25
35
29.5
51.2
90.5
65.8
43.5
9.1
4.7
5.3
0.4
a
From Byrne et al. (1988).
by extending the foot tip, anchoring it with mucus, and
pulling the body forward by pedal muscle contraction
(Wu and Trueman, 1984). Juvenile C. fluminea crawl
on hard subtrates in the same manner, while adults
similarly crawl while lying on one of the valves
(Cleland and McMahon, unpublished). Small specimens of D. polymorpha are active crawlers and can
climb smooth vertical surfaces. They routinely discard
their byssus and migrate to a new position for reattachment. Thus, juveniles migrate to deeper waters in
the winter to avoid ice scour (Mackie et al., 1989).
Freshwater bivalves also respond to external cues.
Chief among these is valve closure in response to irritating external stimuli when detected by mantle edge
and siphonal sense organs (Section II.B.6). Valve closure seals internal tissues from damage by external irritants. Almost all freshwater bivalves tolerate some degree of anaerobiosis (Section II.C.3). Thus, individuals
exposed to irritants may keep the valves shut for relatively long periods until normal external conditions return. Valve closure occurs in response to heavy metals
(Doherty et al., 1987), chlorine, and other biocides
(Mattice et al., 1982; McMahon and Lutey, 1988), suspended solids (Aldridge et al., 1987), and pesticides
(Borcherding, 1992). This ability allows C. fluminea to
avoid intermittent exposure to chlorination or other
biocides, making chemical control of their fouling difficult (Mattice et al., 1982). Valve closure in immediate
response to mantle edge or siphonal tactile stimulation
is a predator defense mechanism. Valve closure response in D. polymorpha is used to monitor water pollution in Europe (Borcherding and Volpers, 1994).
Freshwater bivalves have adaptive behavioral responses to prolonged emersion. Physiological adaptations were discussed in Section II.C.4. Here, behavioral
responses are described in greater detail. When emersed,
C. fluminea displays four major behaviors: (1) escape
responses involving attempts to burrow; (2) valves
gaped widely with mantle edges parted, opening the
mantle cavity to the atmosphere; (3) valves narrowly
gaped with mantle edges exposed, but cemented to-
gether with mucus; and (4) valves clamped shut. Behaviors (1) and (2) are never displayed more than 6% of
time in air. Exposure of sealed mantle edges allows aerial gas exchange (Byrne et al., 1988), but results in
evaporative water loss. When valves are closed, water
loss is minimized; but O2 uptake ceases (Byrne et al.,
1988; McMahon and Williams, 1984). As temperature
increases (Table IV) or relative humidity decreases, duration of mantle edge exposure decreases (McMahon,
1979b; Byrne et al., 1988 ). Hence, behaviors associated with water loss are reduced in response to increased desiccation pressure. Indeed, relative humidities
near zero or temperatures above 30 – 35°C induce continual valve closure (Byrne et al., 1988) (Table IV),
reducing water loss, but making individuals anaerobic.
Similar behaviors occur in emersed unionoideans (Byrne
and McMahon, 1994). Less emersion-tolerant
unionoideans, like P. grandis, display high levels of
mantle edge exposure (25 – 90% of time emersed) and
high rates of water loss. More emersion-tolerant
species, like Uniomerus tetralasmus and Toxolasma
parvus, have reduced levels of mantle edge exposure behavior when emerged (25% of time emersed). The extremely emersion-tolerant U. tetralasmus plugs the
siphon openings with mucus to reduce water loss and
spend little time with mantle edges exposed (Byrne and
McMahon, 1994). Ability of emersed bivalves to adjust
aerial respiratory responses to external dessication pressures is a complex behavior, requiring capacity to sense,
integrate, and respond to external temperature and relative humidity levels and internal osmotic concentration.
Freshwater bivalves may also have circadian patterns of behavior. Both ion-uptake and O2-consumption
rates are greater during dark than light hours in
unionoideans and C. fluminea (McCorkle et al., 1979;
Graves and Dietz, 1980; McCorkle-Shiley, 1982; Section II.C.2) indicative of circadian-activity patterns in
feeding, reproduction, and burrowing. The density of
juvenile C. fluminea in the water column increases during dark hours (McMahon, unpublished) suggesting
that adults preferentially release juveniles at night.
376
Robert F. McMahon and Arthur E. Bogan
Freshwater bivalves may have circadian burrowing cycles, retreating into sediments during light hours to
avoid visual predators like fish and birds. The
unionoideans, Anodonta anatina and Unio tumidis, displayed a diurnal activity rhythm with valves open for
longer periods at night than during the day (Englund
and Heino, 1994a). Similar diurnal valve movement behavior occurs in D. polymorpha (Borcherding, 1992).
2. Filter Feeding
a. Particle Capture Most freshwater bivalves feed
by filtering suspended material from gill water flow
(Section II.B.2) driven by lateral cilia located on each
side of the filaments. Projecting laterally from each filament’s leading edge are “cirri” made up of partially
fused tufts of eulaterofrontal cilia occurring at intervals
of 2 – 3.5 m (Figs. 5A and 21). Eulaterofrontal cirri
on adjacent filaments project toward each other forming a stiffened grid on which suspended seston (phytoplankton, bacteria, and fine detritus) is retained from
water flow driven between the filaments by the lateral
cilia. Filtering may also involve creation of eddies by
cirri in which seston particles settle. Filtered particles
range from 1 to 10 m, with D. polymorpha and C.
fluminea both able to filter smaller particles ( 1.0 m)
than most other freshwater species (Fig. 22) (Jørgensen
et al., 1984; Paterson, 1984; Way et al., 1990a; Silverman et al., 1995). It has been claimed that a mucus net
covering the gill is the primary filter. While the role of
mucus in bivalve filtering is debated, there is little
doubt that the dense mesh of the cirri could form an effective filter (Morton, 1983; Way, 1989). Particles are
captured at the level of the cirri (Jørgensen, 1996), suggesting their involvement in filtration. However, adjacent cirri are separated by 2 – 3.5 m, which until recently, made their mode of particle capture an enigma
because filtered particles were much smaller than the
intercirral distance. Two-particle capture mechanisms
have been hypothesized: direct physical cirral filtration;
or capture by water currents created by gill cilia (Silverman et al., 1996a). Recent confocal microscopic examinations of living gill tissue of D. polymorpha (Silverman et al., 1996a, 1996b) and marine mytilid mussels
(Silverman et al., 1999) support direct cirral particle
capture as the main feeding mode.
The cirrus is composed of a pair of adjacent fused
ciliary sheets aligned with inhalant water flow, extending from a single gill epithelial cell (Fig. 21D.b). Thirtyeight-to-42 cilia form the cirrus. Each cilium has a distinct basal “hinge” region (Fig. 21D.c) and a free distal
end characterized by reduced numbers of microtubules.
Cilia increase in length from the inhalant to the trailing
edge of the cirrus (Fig. 21D.b). Cirral cilia move in unison on the hinge. When they reflex into the space be-
tween adjacent filaments, the free cirral cilia tips splay
out into the inhalant current forming a filter of submicron dimensions (Fig. 21D.a) on which previously freemoving particles (0.75 m diameter) are trapped.
When the cirrus is then flexed back over the leading
edge of the filament, trapped particles are released
(Silverman et al., 1996a, 1996b, 1999) (Fig 21D.a), allowing them to entrain in mucus transported either
ventrally or dorsally by filament frontal cilia (Beninger
et al., 1997) to specialized, ciliated food grooves on the
ventral edges or bases of the demibranches depending
on species (Figs. 2 and 5A – C). In the unionoidean,
Pyganodon cataracta, particles and mucus transported
by filament frontal cilia become entrained and concentrated in a mucus thread on reaching the ventral food
grooves where they are carried anteriorly to the labial
palps (Tankersley, 1996) (Figs. 5A and 21). The role of
mucus in bivalve food particle transport is being
debated Other investigators feel that ciliary-based,
fluid-mechanical forces are primarily responsible for
particle transport with mucus playing a minor role if
any (Jørgensen, 1996). Frontal cirri similar to those of
D. polymorpha occur in sphaeriid, unionoidean and
corbiculacean bivalves (Fig. 21) (Way et al., 1989) and
marine bivalves (Beninger et al., 1997) where they are
presumed to similarly function in particle capture.
b. Sorting of Filtered Particles Mucus-entrained, filtered particles are carried anteriorly to the labial palps,
paired triangular flaps on each side of the mouth (Figs. 2
and 4). The outer labial palp lies against the outer ventral side of the outer demibranch, and the inner palp,
against the inner ventral side of the inner demibranch
(Fig. 2). Palp epithelial surfaces facing away from the
demibranches are smooth, while those adjacent to the
demibranch have a series of parallel ridges lying
obliquely to a ciliated oral groove formed in the fused
dorsal junction of the inner and outer labial palps leading to the mouth (Morton, 1983). In the unionoidean, P.
cataracta, the palps transfer the food-laden mucus
thread from the ctenidial food groove directly to the
mouth for ingestion (Tankersley, 1996). In marine eulammelibranch bivalves, the mucus thread becomes mechanically fluidized on the palp surface by a combination of grinding movements of the opposing palp
surfaces, transport across the corrugated palp surface
and chemical reduction in mucus viscosity. Fluidization,
breaks the mucus thread into a fragmented, flocculent
mucus – particle slurry prior to ingestion (Beninger et al.,
1997). After fluidization, the corrugated palp surface
sorts filtered particles into those accepted for ingestion
and those rejected. Generally, smaller particles are carried over the tops of palp corrugations by cilia to be deposited in the oral groove for ingestion. Denser, larger
11. Mollusca: Bivalvia
FIGURE 21
Scanning electron micrographs of the gill ciliation of representative freshwater bivalves. (A)
Musculium transversum (Sphaeriidae): (a) arrangement of frontal and eulaterofrontal ciliation forming cirri on
the leading edge of the gill filaments in the midgill region; (b) oblique view of a cross-sectional fracture of the
gill showing frontal and eulataterofontal cilia forming cirri on the leading edge of the filament and lateral cilia
on the side of the filament; (c) ciliation of the food groove on the ventral edge of the gill; (d) posterior portion
of the outer (foreground) and inner (background) demibranchs of the gill; (e) a frontal cirrus formed from
fused cilia emerging between the eulaterofrontal cirri and frontal cilia on the leading edge of a filament; (f) a
frontal cirrus emerging from a band of frontal cilia. (B) Corbicula fluminea (Corbiculidae): (a) leading edge of
a gill filament showing frontal cilia, eluatorofrontal cilia forming cirri, lateral cilia and frontal cirri; (b) highmagnification view of frontal cilia and eulateralfrontal cirri; (c) oblique view of a longitudinal fracture through
the midgill region showing all four types of ciliation, including lateral cilia lining the sides of the gill filiament;
(Continues)
377
378
Robert F. McMahon and Arthur E. Bogan
FIGURE 21
(Continued)
(d) crosssection of the midgill region showing the origin of the frontal cirrus; (e) lower region of the gill showing the presence of less well organized frontal cilia; (f) food groove region at the ventral edge of the inner demibranch; note loss of ciliary organization in food groove, including lack of frontal cirri. (C) Obliquaria reflexa
(Unionoidea): (a) frontal cilia and eulaterofrontal cilia forming cirri on the leading edges of several gill filaments (500X); (b) high-power view showing lateral cilia on sides of the gill filaments between tracts of frontal
cilia and eulaterofrontal cilia forming cirri on adjacent filaments. (D) Dreissena polymorpha (Dreissenoidea):
(a) frontal cilia on the leading edge of a gill filament and some eulaterofrontal cirri reflexed into the space
between adjacent filaments with the free, distal ends of their cilia splayed out to form the primary gill filtering
mechanism and other cirri reflexed over the frontal cilia in a position in which they release particles captured
from water flow between adjacent filaments to frontal cilia of the food groove; (b) detailed structure of a eulaterofrontal cirrus formed from two rows of opposed cilia fused basally, free at their distal tips, with a distinct
(Continues)
11. Mollusca: Bivalvia
FIGURE 21
(Continued)
basal hinge on which the cirral cilia simultaneously flex between the interfilament space (food capture) and the
frontal cilia on the leading edge of the filament (food release); (c) transmission electron micrograph showing
details of the unique hinge on which cirral cilia flex between the interfilament space and the frontal cilia. Note
lack of frontal cirri characteristic of the (A) Sphaeriidae, and (B) Corbicula fluminea in (C) unionoideans and
(D) Dreissena Label key for figures A – C : F, frontal cilia; FC, frontal cirrus; L, lateral cilia, LF eulaterofrontal
cirri; and FG, food groove. [Photomicrographs supplied by Tony Deneka and Daniel Hornbach (Macalester
(Continues)
379
380
Robert F. McMahon and Arthur E. Bogan
FIGURE 21
(Continued)
College), Carl M. Way (U.S. Army Corps of Engineers), and Harold Silverman and Thomas H. Dietz
(Louisiana State University, Baton Rouge).]
11. Mollusca: Bivalvia
particles fall between the corrugations where cilia carry
them to the ventral edge of the palp to be bound in mucus and released onto the mantle as pseudofeces (pseudofeces are mucus-bound particles that have not been
ingested) (Morton, 1983; Beninger et al., 1997). Pseudofeces are carried by mantle cilia to the base of the inhalant siphon (Fig. 2), where they are periodically expelled with water forced from the inhalant siphon by
rapid valve adduction (i.e., valve clapping) (Morton,
1983). The palp may also sort medium-sized particles.
Smaller medium-sized particles tend to remain longer on
corrugation crests than in intervening grooves, thus,
reaching the oral groove for ingestion, while larger,
medium-sized particles remain longer in the grooves
leading to eventual rejection as pseudofeces. Thus, labial
palp particle sorting may be primarily by particle size
and, perhaps, density. The function of labial palps in
particle sorting is being debated, palps appearing to have
little if any sorting function in some species (Tankersley,
1996) and extensive sorting functions in others
(Beninger et al., 1997). Labial palp function in freshwater bivalves requires further study.
The width of palp corrugations and gill filaments
can be adjusted by both muscular activity and distention of gill and palp blood sinuses, thus allowing active
particle-size selection (Morton, 1983). When fed
natural seston, the percentage of particles retained by
the unionoidean, Elliptio complanata, declined linearly
over a particle size range of 5 (nearly 100%) to
10m (less than 10% retention), suggesting size selection (Paterson, 1984). Similar size selection for particles occurs in the sphaeriid, Musculium transversum
(Way, 1989), and C. fluminea (Way et al., 1990a).
Some unionoidean species may select different algal
types. In the same river pool, Amblema plicata ingested
a greater proportion of green algae, a lower diversity of
algal species, and different algal species than Ligumia
recta (Bisbee, 1984). Interspecific differences in particle
selection may allow some sympatric species to divide
the feeding niche, avoiding competition, however, other
unionoideans are nonselective in feeding and have high
diet overlap between species.
c. Filtering Rate The filtering rate of bivalves can
be computed as follows:
冢
冣
C0
M
In
t
Ct
where M is the volume of suspension filtered, t the
time, and Co and Ct the concentrations of filtered particles at time o and t, respectively (Jørgensen et al.,
1984). Filtration rates are elevated in C. fluminea
compared to other freshwater bivalves. For a 25-mm
shell length (SL) specimen, rate ranges from 300 to
C
381
2500 L / h depending on temperature and seston
concentration (Foe and Knight, 1986b; Lauritsen,
1986b; Way et al., 1990a). The filtering rate of C. fluminea fed natural seston concentrations was not significantly correlated with field water temperature (Long
and McMahon, unpublished) and was independent of
temperature between 20 and 30°C in the laboratory
(Lauritsen, 1986b), suggesting capacity for temperature
compensation of gill ciliary filtering activity. The filtration rate is also somewhat temperature independent in
Sphaerium striatinum and Musculium partumeium,
reaching maximal values during reproductive periods
when water temperatures, were below midsummer
maxima (Burky et al., 1981, 1985a; Way et al., 1981).
Filtering rates in D. polymorpha are similar to those in
C. fluminea (Lei et al., 1996) and 2 – 4 times higher
than those in sphaeriids and unionoideans (Kryger and
Riisgård, 1988).
High particle concentrations reduce filtering rates
(Burky et al., 1985a; Way, 1989; Way et al., 1990a; Lei et
al., 1996). Filtering rates in S. striatinum (Hornbach
et al., 1984a) and M. partumeium (Burky et al., 1985a)
were depressed at concentrations below natural seston,
making particle ingestion nearly constant above a critical
particle concentration. Thus, filtering may have be regulated to control ingestion rate. Particle concentrations
within ambient seston concentrations do not affect filtering rate in C. fluminea (Mattice, 1979), ingestion rates
increasing directly with concentration. However, filtration rate in C. fluminea declined three-fold with increasing particle concentration between 0.33 and 2.67 l algal
volume / L suggesting inhibition at very high concentrations, but increasing algal concentration over this range
still resulted in a 3.5-fold rise in ingestion rate (Lauritsen,
1986b). Similar reduction in filtration rate at particle
concentrations above natural seston levels occurs in the
sphaeriid, M. transversum (Way, 1989). In contrast to C.
fluminea, filtration rate of the unionoidean, E. complanta, was depressed by increasing natural seston concentrations (Paterson, 1984). Filtration rate differed
among populations of C. fluminea, but ingestion rates
were similar, suggesting regulation of the filtration rate to
control ingestion rates (Way et al., 1990a). In D. polymorpha, filtration rate was reduced when feeding on artificial plastic microspheres compared to natural seston.
When filtering particles 1.5 m in diameter, filtration
rate decreased with decreasing particle concentration below a critical upper limit, and with decreasing temperature. Filtration rate was also affected by prior temperature acclimation (Lei et al., 1996). Such data indicate that
the seasonal and environmental filter-feeding responses of
freshwater bivalves are species-specific and laboratory
measurements of filtration rates are likely to be invalid
unless carried out at natural seston concentrations and
382
Robert F. McMahon and Arthur E. Bogan
sizes and with natural seston rather than artificial particles. Filter feeding and particle retention rates were depressed in gravid females of the unionoidean, Pyganodon
cataracta, in which particle transport was slowed down
and the ability to retain small particles (6 m) reduced,
when the marsupial gills were distended with glochidia
(Tankersley, 1996).
Suspended silt can inhibit filtering and consumption rates in freshwater bivalves, perhaps by overwhelming ciliary filtering and sorting mechanisms.
High silt concentrations reduced filtering and metabolic
rates in three unionoidean species whether exposure
was infrequent (once every 3 h) or frequent (once every
0.5 h), suggesting interference with gill ciliary activity.
Furthermore, individuals experiencing frequent exposure increased reliance on carbohydrate catabolism
(Aldridge et al., 1987), symptomatic of short-term starvation; perhaps, the basis of exclusion of unionoideans
from habitats with high silt loads (Adam, 1986). In
contrast, natural suspended sediment concentrations
did not affect shell or tissue growth in C. fluminea (Foe
and Knight, 1985), perhaps, accounting for its ability
to colonize high-flow lotic habitats with greater suspended solids than tolerated by most unionoideans
(Payne and Miller, 1987; Section III.A.). In contrast,
growth in juvenile unionoideans is stimulated by small
amounts of suspended silt in their algal food (Hudson
and Isom, 1984), perhaps because juveniles (Villosa
iris) may feed primarily on sediment interstitial water
through the pedal gape, leading to ingestion of silt,
microdetritus and bacteria, with little consumption of
algae (Yeager et al., 1994).
In smaller streams, where phytoplankton productivity may be low, most suspended organic matter is
particulate organic detritus or heterotrophic bacteria
and fungi (Nelson and Scott, 1962). While experiments
are few, at least some freshwater bivalves efficiently
filter suspended bacteria. D. polymorpha appears to
extract particles of bacteria size (1.0 m in diameter)
(Sprung and Rose, 1988, Fig. 22) as can C. fluminea
(Silvermann et al., 1995) with D. polymorpha and C.
fuminea able to clear the bacterium, Escherichia coli,
30 and 3 times faster than the lotic unoinid, Toxolasma
texasiensis, respectively. Increased capacity for bacterial filtration in D. polymorpha appears associated with
its enlarged gill with a 100-fold greater density of cirri
relative to C. fluminea, that of T. texasiensis has even
fewer and smaller cirri than D. polymorpha or C. fluminea (Silverman et al., 1995). Filter-feeding by D.
polymorpha may significantly reduce bacterioplankton
densities in the inner portion of Saginaw Bay, Lake
Huron (Cotner et al.,). Musculium transversum (Way,
1989a) and some unionoidean species (Fig. 22) do not
efficiently filter bacterial-size particles. Unionoideans
appear specialized to filter larger particles, such as phy-
FIGURE 22
Relationship between the percentage of suspended
particles retained by the gill filtering mechanism of freshwater
bivalves and particle size, as particle diameter in micrometers. Horizontal axis for all figures is the percentage of total particles retained
from suspensions passing over clam gill filtering systems in (A) Dreissena polymorpha, the unionoideans, (B) Unio pictorum, and (C) Anodonta cygnea. Vertical lines about points are standard deviations.
Note that all three species efficiently retained particles of 2.5 – 8 m
diameter, equivalent to most unicellular algae, but only D. polymorpha efficiently retained bacteria-sized particles of 1 m diameter.
(Redrawn from Jørgensen et al., 1984.)
toplankton, detritus, protozoa, and even small zooplankton, such as rotifers (Singh et al., 1991). In contrast, six unionoidean species adapted to lotic habitats
where seston was dominated by bacteria and detritus
could filter bacteria at significantly higher rates than
three species adapted to lentic habitats where seston
was dominated by phytoplankton. Lotic species had
more complex cirri (25 vs 16 cilia per cirral plate)
and greater gill cirral density than lentic species, indicative of a filtering system better adapted for fine particle
filtration (Silverman et al., 1997) . Because bacteria and
detrital particles may comprise most of the organic
matter in some, particularly lotic, habitats, bacterial
feeding warrants greater study in freshwater bivalves.
d. Sediment Feeding At least some freshwater bivalves may supplement phytoplankton filter feeding by
consuming organic detritus or interstitial bacteria from
sediments. The infaunal habit of many Pisidium species
suggests dependence on nonplanktonic food sources.
Many pisidiids can efficiently filter small interstitial bacteria. They burrow continually through sediments with
the shell hinge downward, drawing dense interstitial
11. Mollusca: Bivalvia
water and bacteria through parted ventral mantle edges
into the mantle cavity to be filtered on the gills (Lopez
and Holopainen, 1987). Both Pisidium casertanum and
P. conventus feed in this manner, filtering bacteria of
smaller diameter (1 m) than most bivalves (Lopez
and Holopainen, 1987) (Fig. 22). Filtering of rich interstitial bacterial flora may be associated with the reduced
ctenidial surface areas and low filtration and metabolic
rates of Pisidium species compared to Sphaerium or
Musculium (Hornbach, 1985; Lopez and Holopainen,
1987). Similar interstitial bacteria and detritus feeding
occurs in juvenile unionids (Yeager et al., 1994). M.
transversum uses its long inhalant siphon to vacuum detrital particles from the sediment surface (Way, 1989).
Pedal feeding on sediment organic detritus may be
another important food source in some freshwater bivalves. Pedal feeding occurs in marine and estuarine bivalves (Morton, 1983; Reid et al., 1992), but is little
studied in freshwater species. Corbicula fluminea draws
sediment detrital particles over the ciliated foot epithelium into the mantle cavity, where they accumulate in
the food groove of the inner demibranch and are carried
to the palps for sorting and ingestion (Way et al.,1990b;
Reid et al., 1992). Cellulose particles (40 – 120 m in diameter) soaked in various amino acid solutions and
mixed with sediments accumulated in the stomach of
pedal-feeding C. fluminea (McMahon, unpublished).
Thus, C. fluminea removed sediment organic matter at a
rate of 50 mg / clam per day and doubled growth rates
when allowed to pedal feed in addition to filter feeding
(Hakenkamp and Palmer, 1999). Similar pedal feeding
has been reported in M. transversum (Way, 1989) and in
juvenile Villosa iris (Yeager et al., 1994). Thus, pedal deposit feeding may be an important auxiliary feeding
mode in freshwater and marine bivalves and may have
important impacts on sediment organic dynamics
(Hakenkamp and Palmer, 1999).
Pedal feeding may be more universal in freshwater
species than previously suspected. In a stream S. striatinum population, only 35% of organic carbon assimilation occurred by filter feeding, leaving 65% to come
from sediment detrital sources (Hornbach et al.,
1984b), presumably by pedal feeding. Other sphaeriid
species reach highest densities in sediments of high organic content (Zbeda and White, 1985) or in habitats
receiving organic sewage effluents, suggesting dependence on pedal deposit-feeding (Burky, 1983). Pedal
deposit-feeding mechanisms may explain the extensive
horizontal sediment locomotion of a number of freshwater bivalves (Section II.D), as it allows pedal contact
with organic detritus.
3. Population Regulation
a. Abiotic Population Regulation Most of the information regarding regulation of freshwater bivalve
383
populations is anecdotal. Fuller (1974) reviews factors
affecting population density and reproductive success.
Catastrophic abiotic factors causing reductions in bivalve populations are reviewed in Section III.B. Among
these are silt accumulation in sediments of impounded
rivers and in the water column during flooding. Silt interferes with gill filter feeding and gas exchange
(Aldridge et al., 1987; Payne and Miller, 1987) causing
massive mortality in unionoidean populations (Adam,
1986, Section III.A). In contrast, many sphaeriids thrive
in silty sediments (Burky, 1983; Kilgour and Mackie,
1988) associated with their utilization of sediment detrital food sources (Section III.C.2). Shell growth in C.
fluminea, a silt tolerant species, can be temporarily
inhibited by high turbidity (Fritz and Luz, 1986).
Temperature extremes also affect bivalve populations. C. fluminea tolerates 2°C (Mattice, 1979) to
36°C (McMahon and Williams, 1986b). Both cold-induced winter (Blye et al., 1985; Sickel, 1986) and heatinduced summer kills (McMahon and Williams,
1986b) are reported for this species. The incipient upper lethal limit for D. polymorpha is 30°C, preventing
its invadsion of many southern U.S. drainage systems
(McMahon, 1996). Temperature limits for other NA
bivalves are not as well known, but, on average, appear to be broader than either C. fluminea or D. polymorpha (Burky, 1983). Juvenile unionoideans are less
temperature tolerant than adults (Dimock and Wright,
1993), suggesting that temperature limitation operates
at the juvenile level in this group. Within the tolerated
range, temperature may have detrimental effects on reproductive success. Sudden temperature decreases
cause abortive glochidial release in unionoideans
(Fuller, 1974), and temperatures above 30 – 33°C inhibit reproduction in C. fluminea (McMahon, 1999).
Sphaeriid densities were reduced in midsummer by
heated effluents (Winnell and Jude, 1982). Heated effluents may also stimulate bivalve growth, inducing
early maturity and increasing reproductive effort. They
can provide heated refugia in which cold-sensitive
species may overwinter as in northernmost NA populations of C. fluminea (McMahon, 1999).
Low ambient pH can depauperate or extirpate bivalve populations. Bivalve populations have been extirpated by acid mine drainage (Taylor, 1985; Warren
et al., 1984). Highly acidic waters cause shell erosion
and eventual death (Kat, 1982). Perhaps of more importance than pH in regulating freshwater bivalve
populations are water hardness and alkalinity.
Unionoideans do not occur in New York drainage systems with calcium concentrations 8.4 mg Ca / L
(Fuller, 1974). In waters of low alkalinity, Ca concentrations may be too low for shell deposition; the lower
limit for most freshwater bivalves being 2 – 2.5 mg
Ca / L (Okland and Kuiper, 1982; Rooke and Mackie,
384
Robert F. McMahon and Arthur E. Bogan
1984a). Growth and fecundity were suppressed in a
Pisidium casertanum population in a low Ca concentration relative to that a higher Ca concentration
(Hornbach and Cox, 1987). Waters of low alkalinity
have little pH-buffering capacity, subjecting them to
seasonal pH variation. This makes them particularly
sensitive to acid rain, which detrimentally impacts bivalve faunae (Rooke and Mackie, 1984a). Shell Ca
content is related to water Ca concentration. Of 10
Canadian freshwater bivalve species, two sphaeriids
and one unionoidean displayed no relationship between shell Ca content and water Ca concentration,
shell Ca content decreased with increased water Ca
concentration in two sphaeriid species and increased
with water Ca concentration in four sphaeriid and two
unionoidean species (Mackie and Flippance, 1983b).
Lowering of water levels during dry periods induce
bivalve mortality by exposing individuals to air. Such
emersion caused near 100% mortality in C. fluminea
populations (White, 1979) due to its poor desiccation
tolerance (Bryne et al., 1988). Many unionoidean
species are relatively tolerant of emersion. When sympatric populations of nine unionoidean species, the
sphaeriid Musculium transversum, and C. fluminea
were emersed for several months, M. transversum and
C. fluminea suffered 100% mortality while the
unionoideans experienced only 50% mortality, because
they either migrated downshore or resisted desiccation
(White, 1979). Like C. fluminea, D. polymorpha is relatively emersion intolerant, particularly above 20°C
(McMahon, 1996), restricting it to habitats with relatively stable water levels and making it unlikely to develop dense populations in variable level reservoirs.
Low environmental O2 concentrations can be detrimental to freshwater bivalves. Continual eutrophication of Lake Estrom, Denmark, so reduced ambient
profundal O2 concentrations (0 – 0.2 mg O2 /L for three
months) that massive reductions of sphaeriid populations ensued (Jonasson, 1984a, b), including hypoxiatolerant species, such as Pisidium casertanum and P.
subtruncatum in which the lower limit for aerobic respiration was 1.7 mg O2 /L (Jonasson, 1984b). Adult
unionoideans are relatively hypoxia tolerant. Juveniles
are more hypoxia sensitive, those of Utterbackia imbecillis and Pyganodon cataracta tolerating anoxia for
less than 24 h (Dimock and Wright, 1993). Thus, restriction of unionoidean species to well oxygenated waters may be a juvenile rather than adult constraint.
Pollution is also detrimental to freshwater bivalves
(reviewed by Fuller, 1974) including chemical wastes
(Zeto et al.,1987), asbestos (Belanger et al.,1986a), organic sewage effluents (Gunning and Sutkkus, 1985;
Neves and Zale, 1982; St, John, 1982), heavy metals
(Belanger et al., 1986b; Fuller, 1974; Lomte and
Jabhav, 1982a; Grapentine, 1992), chlorine and paper
mill effluents (Fuller, 1974), acid mine drainage (Taylor,
1985; Warren et al., 1984) and PCBs (Grapentine,
1992). Potassium ions, even in low concentration
(4 – 7 mg K /liter), can be lethal to freshwater bivalves including D. polymorpha (Claudi and Mackie,
1993), excluding bivalves from watersheds where
potassium is naturally abundant (Fuller, 1974).
b. Biotic Population Regulation There have been a
few studies of biotic regulation of freshwater bivalve
populations. Freshwater bivalves are host for a number
of parasites. They are intermediate hosts for digenetic
trematodes (Fuller, 1974). Such infections cause sterility
in gastropods, and have been reported to induce sterility
in the unionoidean, Anodonta anatina, allowing diversion of the mussel’s energy stores from production of its
gametes to production of trematode cercariae (Jokela et
al., 1993), reducing the mussel population’s reproductive capacity. Parasitic nematode worms inhabit the guts
of unionoideans (Fuller, 1974). The external oligochaete
parasite Chaetogaster limnaei resides in the mantle cavities of unionoideans (Fuller, 1974), C. fluminea (Sickel,
1986) and D. polymorpha (Conn et al., 1996). Dreissena polymorpha is host to a wide diversity of parasites
(Malloy et al., 1997), but their impacts on this species
are unknown. All of these parasites probably contribute
to the regulation of freshwater bivalve population densities, but the degree to which they do has received little
experimental attention.
Water mites of the family Unionicolidae, including
Unionicola and Najadicola, are external parasites of
unionoideans (Vidrine, 1990). Both mature and preadult mites are parasitic, attaching to gills, mantle, and
the visceral epithelium (depending on species) (Mitchell,
1955; Chapter 16). Heavy mite infestations cause
portions of the gills to be shed, abortion of developing
glochidia, or even death (Fuller, 1974), perhaps making
unionicolid mites a major regulator of unionoidean
populations. The larval stage of the chironomid,
Ablabesmyia janta, parasitizes the unionoidean gill filaments and may exclude unionicolid mites from the gills
they infest (Vidrine, 1990).
Disease has been little studied. There is little evidence of viral or bacterial involvement in die-offs of C.
fluminea (Sickel, 1986) or unionoideans (Fuller, 1974).
In some C. fluminea populations, massive die-offs
(particularly of older individuals) occur after reproductive efforts (Aldridge and McMahon, 1978; McMahon
and Williams, 1986a; Williams and McMahon, 1986),
probably due to reductions in tissue energy reserves of
postreproductive individuals (Williams and McMahon,
1989). Such postreproductive morality also occurs in
sphaeriids (Burky, 1983).
11. Mollusca: Bivalvia
TABLE V List of the Major Fish Predators
of Freshwater Bivalvesa
Family
Genus and species
Common name
Clupeidae
Cyprinidae
Catostomidae
Alosa sapidissima
Cyrinus carpio
Ictiobus bubalus
Ictiobus niger
Minytrema melanops
Moxostoma carinatum
Morone saxatilis
Ictalurus furcatus
Ictalurus punctatus
Lepomis gulosus
Lepomis macrochirus
Lepomis microlophus
Aplodinotus grunniens
Acipenser fulvesens
American shad
Common carp
Smallmouth buffalo
Black buffalo
Spotted sucker
River redhorse
Striped bass
Blue catfish
Channel catfish
Warmouth
Bluegill
Red-ear sunfish
Freshwater drum
Lake sturgeon
Moronide
Ictaluridae
Centrarchidae
Sciaenidae
Acipenseridae
a
Data from Fuller (1974), McMahon (1983a), Robinson and
Wellborn (1988), and Sickel (1986).
Predation may be the most important regulator of
freshwater bivalve populations. Shore birds and ducks
feed on sphaeriids and C. fluminea (Dreier, 1977;
Paloumpis and Starrett, 1960; Smith et al., 1986;
Thompson and Sparks, 1977, 1978) and D. polymorpha (Mazak et al., 1997). C. fluminea densities were
3 – 5 times greater in enclosures excluding diving ducks
(Smith et al., 1986). Water fowl feeding also significantly reduces D. polymorpha populations (Mackie
and Schloesser, 1996). Crayfish feed on small bivalves
including C. fluminea (Covich et al.,1981) and D. polymorpha (Mackie et al., 1989), and, in some habitats,
may regulate bivalve population densities. Fire ants
(Solenopis invicta) prey on clams emersed by receding
water levels. In additions, turtles, frogs, and the mudpuppy salamander Necturus maculosus all feed to a
limited extent on small or juvenile bivalves (Fuller,
1974). Free-living oligochaetes prey on newly released
glochidia (Fuller, 1974).
Perhaps the major predator of freshwater bivalves
are fish. A number of NA fish are molluscivores
(Table V). While most molluscivorous fish feed on small
bivalve species or juveniles of larger bivalves
(SL 7 mm), several routinely take large adults, including carp (Cyprinus carpio), channel catfish (Ictalurus
punctatus) and freshwater drum (Aplodinotus grunniens). The latter species crushes shells with three
massive, highly muscular pharyngeal plates, as do carp
to a lesser extent. Bivalve prey size may be limited by
the dimensions of the shell-crushing apparatus. Thus,
273 – 542-mm long A. grunniens did not take D. polymorpha with SL 11.4 mm, but, below this size, fish
size and mussel size taken were not related. Thus,
385
predation by A. grunniens is unlikely to control D.
polymorpha populations (French and Love, 1995). In
contrast, channel catfish swallow bivalves intact (J. C.
Britton, personal communication). The majority of molluscivorous fish may limit predation to smaller bivalves
because shells of larger individuals are too strong to
crack or crush. The shell strength of C. fluminea increases exponentially with size [log10 force in newtons
to crack the shell 0.76 2.31 (log10 SL in mm)]; it
is 6.5 times stronger than the very thick-shelled estuarine bivalve, Rangia cuneata (Fig. 23) (Kennedy and
Blundon, 1983). Shell strength is directly correlated
with shell thickness. In specimens with an SL of 20 mm,
the 3.0 mm thick shell of the unionoidean, Fusconaia
ebena, was more crush resistant than the 1.3 mm thick
shell of C. fluminea which, in turn, was more crush resistant than the fragile 0.31 mm thick shell of D. polymorpha. The fragile shell of D. polymorpha makes it
prone to predation by diving ducks, crayfish and fish
relative other freshwater bivalves (Miller et al., 1994).
Fish predation can deplete freshwater bivalve populations. Sphaeriid diversity and density increased in
habitats from which molluscivorous fish were excluded
(Dyduch-Falniowska, 1982). Robinson and Wellborn
(1988) reported that, 11 months after settlement, densities of C. fluminea were 29 times greater in enclosures
excluding fish. In contrast, exclusion of fish and turtles
did not increase density or species richness of a combined unoinoidean and sphaeriid community in a cooling reservoir (Thorp and Bergey, 1981).
As fish are intermediate hosts for unionoidean
glochidia, the size of fish host population can influence
mussel reproductive success. Absence of appropriate
fish hosts have caused extirpation of unionoideans in a
number of NA aquatic habitats (Fuller, 1974; Kat and
Davis, 1984; Smith, 1985a, b; Neves et al., 1997;
Vaughn, 1997; Section III. A.) and availability of fish
hosts may regulate recruitment (Johnson and Brown,
1998). Indeed, restoration of host fish populations produced remarkable recoveries of some endangered
unionoidean populations (Smith, 1985b). Thus, anthropomorphic activities reducing fish host population densities can result in destruction of unionoidean populations, making relationships between unionoideans and
their glochidial host fish an important future management consideration for drainage systems and their fisheries to ensure continued health of their remaining
unionoidean fauna (Watters, 1993; Neves, 1997).
Mammals prey on freshwater bivalves. Otters,
minks, muskrats, and raccoons eat bivalves and may
regulate populations of some species (Fuller, 1974).
Raccoons and muskrats feed on C. fluminea and may
account for reductions of adult clam densities in the
shallow, near-shore waters of some Texas rivers, where
386
Robert F. McMahon and Arthur E. Bogan
FIGURE 23 Shell strength of Corbicula fluminea. Solid line is the
best fit of a geometric mean estimate of a least squares log–log linear
regression, relating shell length (SL) to force required to crack the
shell (open circles) as follows: Log10 force to crack the shell in newtons 0.76 2.31 (log10 mm SL). Dashed line is the best fit of a
similar log–log linear regression for the estuarine wedge calm, Rangia
cuneata, as follows: log10 force to crack the shell in newtons
0.736 2.42 (log10 SL in mm). R. cuneata, with the strongest
shell of eight tested estuarine bivalve species, has a thicker shell than
does C. fluminea. However, as the regression slope values for the two
species are nearly equal, the elevated y-intercept for C. fluinea
(0.76 compared to 1.73 for R. cuneata) indicates that its shell is
approximately an order of magnitude stronger. (Redrawn from
Kennedy and Blundon, 1983.)
feeding sites are marked by thousands of opened shells
(McMahon, unpublished observations). Even wild hogs
root out and destroy shallow-water unionoidean beds
(Fuller, 1974). Muskrats annually consummed 3% of
individuals of Pyganodon grandis in a small Canadian
lake, equivalent to 31% of the mussel’s annual tissue
production. They preferentially consumed larger mussels (shell length 55 mm), strongly affecting mussel
population size – age structure, resulting in a biomass
decline of large individuals, reducing the population’s
annual reproductive effort (Hanson et al., 1989).
c. Competitive Interactions There have been no experimental evaluations of interspecific or intraspecific
competition among freshwater bivalves; only anecdotal
observations. Unionoidean or sphaeriid population declines coincident with establishment of C. fluminea
have occurred in NA habitats (McMahon, 1999;
Sickel, 1986). C. fluminea may detrimentally impact
sphaeriid populations after invasion, but there is little
evidence of major impacts of C. fluminea on NA
unionoideans (Strayer, 1999b). Thus, C. fluminea had
little impact on a Tennessee River unionoidean community, the C. fuminea population declining over time
while unionid density remained stable (Miller et al.,
1992). Rather, habitats are first made inhospitable to
indigenous unionoideans by channelization, dredging,
over-fishing or other water management practices. C.
fluminea is more tolerant of high flows, suspended
solids, and silting caused by human activities in
drainage systems, allowing it to rapidly colonize such
disturbed habitants in which unionoideans have been
reduced or extirpated (McMahon, 1999). In contrast,
establishment of C. fluminea in drainage systems receiving minimal human interference has little effect on
native bivalve populations (McMahon, 1999; Sickel,
1986), suggesting an inability of Corbicula to outcompete native unionoideans in undisturbed habitats. The
impacts of C. fluminea on native, NA bivalves remains
subject to debate (reviewed by Strayer, 1999b) and
should be experimentally addressed.
In contrast to C. fluminea, there is clear evidence
of direct negative impacts of D. polymorpha on NA
unionoidean populations. Dreissena polymorpha
byssally attaches to the posterior portions of unionoidean valves projecting above the substrate where
they can reach densities which strip seston from the
unionoidean’s inhalant current, leading to its slow
starvation. Extensive D. polymorpha infestations can
cause unionoideans to be dislodged from the substrate,
leading to death. D. polymorpha infestation has
caused near complete extirpation of unionoideans
from the lower Great Lakes and St. Lawrence River
(Schloesser et al., 1996; Strayer, 1999b). Interestingly,
short-term brooding unionoideans (tribes Amblemini
and Pleurobemini) appear less sensitive to zebra mussel impacts than long-term brooders (tribe Lampsilini
and subfamily Anodontinae), perhaps due to their
greater tissue energy reserves (Strayer, 1999b).
Unionoidean populations in shallow (1 m), near-shore
waters of western Lake Erie were less subject to D.
polymorpha infestation than those at greater depths
(Schloesser et al., 1997), perhaps because wave-induced agitation inhibits byssal attachment (Clarke and
McMahon, 1996b).
Dreissena polymorpha can have indirect impacts
on N.A. bivalve communities. After it colonized the
lower Hudson River in 1992, sphaeriid and
unionoidean populations declined even though D.
polymorpha did not attach to their shells. This decline
resulted from a massive reduction in phytoplankton
11. Mollusca: Bivalvia
density by D. polymorpha feeding, causing starvation
of indigenous bivalves (Strayer, 1999b).
d. Density-Dependent Mechanisms of Population
Regulation Factors directly impacting freshwater bivalve population densities may have secondary impacts
associated with reduction in gamete fertilization
through reduction of sperm concentration in the water
column. Thus, egg fertilization in the marsupial gills of
female Elliptio complanata was directly correlated with
population density, fertilization failing completely at
10 adults / m2 (Downing et al., 1993).
Published experimental investigations of intraspecific, density-dependent, population regulation mechanisms in freshwater bivalves are also lacking. However,
some evidence is available from descriptive field studies
of C. fluminea and sphaeriids. In C. fluminea, extremely successful recruitment of newly settled juveniles occurred after adult population decimation
(McMahon and Williams, 1986a, b); whereas, juvenile
settlement is generally far less successful in habitats
harboring dense adult populations. High adult density
may prevent successful juvenile recruitment, thus preventing extensive juvenile – adult competition. This hypothesis was tested by enclosing adult C. fluminea at
different densities in a lake harboring a dense Corbicula population. While adult density had no effects on
adult or juvenile growth rates, it affected juvenile settlement. Settlement in enclosures without adults was
8702 juveniles / m2. Maximal settlement (26,996 juveniles / m2) occurred at 329 adults / m2; and was least at
the highest adult densities of 659 and 1976 clams / m2,
being only 8642 and 3827 juveniles / m2, respectively
(McMahon, unpublished data). Thus, adults of C. fluminea may inhibit juvenile settlement. Mackie et al.,
(1978) found that high adult densities in Musculium securis caused significant reduction in the number of juveniles incubated, implying that intraspecific adult
competition regulates reproductive effort. In addition,
interspecific competition between M. securis and M.
transversum caused reduction in reproductive capacity
in the subdominant species, with species dominance dependent on habitat. While intraspecific density effects
and interspecific competition may regulate Corbicula
and sphaeriid populations, results may not be the same
for highly K-selected unionoidean populations which
require further study before the effects of thinning of
mussel beds by commercial harvesters on juvenile recruitment and adult growth can be assessed.
4. Functional Role in the Ecosystem
Bivalves play important roles in freshwater ecosystems (reviewed by Strayer et al., 1999a, b). Because they
can achieve high densities and are filter feeders, they are
387
important consumers of phytoplankton primary productivity. Unionoideans, with filtering rates on the order
of 300 mL / g dry tissue/ h (Paterson, 1984), can account
for a large proportion of total consumption of phytoplankton productivity. As unionoidean densities range
from 15 individuals / m2 (Paterson, 1985) to 28 individuals / m2 (Negus, 1966) and dry tissue standing crop
biomasses from 2.14 g / m2 (Paterson, 1985) to 17.07
g / m2 (Negus, 1966), filtering by unionoidean communities could be as high as 15 – 122.9 L / m2 per day. Thus,
on an annual basis, a eutrophic lake unionoidean community filtered 79% of total lake volume, removing
92 – 100% of all suspended seston. However, even with
such high cropping rates, high phytoplankton reproductive rates resulted in unionoideans reducing seston by
only 0.44% (Kasprzak, 1986). While the unionoidean
community accounted for only 0.46% of seston removed by all phytoplanktivorous animals in the lake,
they made up 85% of its standing crop biomass. Their
biomass contained a high proportion of the phosphorus
load of the lake, limiting the quantity of phosphorus
available for phytoplankton production. Similarly, the
unionoidean community of Lake St. Clair ingested
13.5% of the total phosphorus load of the lake from
May through October and deposited 64% of consumed
phosphorus in the sediments, providing nutrients for
rooted macrophytic vegetation and invertebrate deposit
feeders (Nalepa et al., 1991). D. polymorphia populations in five European lakes had an average dry weight
biomass 38 times that of submerged macrophytes and
average standing crop phosphorus and nitrogen contents 2.94 (range, 0.39 – 7) and 4.21 (range, 0.59 – 9.4)
times that of submerged macrophytes, respectively,
greatly reducing availability of these growth-limiting
inorganic nutrients to the aquatic plant community
(Stanczykowska, 1984). In the western basin of Lake
Erie, D. polymorpha has become a major factor in
phosphorus flux; its uptake of phosphorus and nitrogen
potentially limiting phytoplankton growth and its low
N:P excretion ratios (20:1) potentially shifting phytoplankton community structure toward cyanobacteria
(Arnott and Vanni, 1996).
Sphaeriids also have important impacts on lentic
phytoplankton communities. The sphaeriid community
of an oligotrophic lake comprised only 0.12% of total
planktivore biomass and 0.2% of the bivalve biomass,
but accounted for 51% of total seston consumption
(Kasprzak, 1986). This probably resulted from their
relatively high filtration rates and population productivity (Burky et al., 1985a; Hornbach et al., 1984a, b).
In smaller lotic and lentic habitats, sphaeriids can consume a major portion of primary productivity. In a
canal community dominated by sphaeriids (98% of
planktivore biomass), they accounted for 96% of total
388
Robert F. McMahon and Arthur E. Bogan
seston consumption (Krasprzak, 1986). A stream population of S. straitinum was estimated to filter 3.67 g organic C / m2 per year or 0.0004% of seston organic carbon flowing over them (Hornbach et al., 1984b). In
contrast, filtration by a pond population of M. partumeium removed 13.8 g C / m2 per year as seston
(Burky et al., 1985a). Based on the data of Burky et al.,
(1985b), standing crop seston levels for the entire pond
ranged annually between 46 and 550 g carbon. Average annual seston organic carbon consumption by the
M. partumeium community was estimated to be
roughly 3.9 g C per day, or 0.7 – 8.5% of total standing
crop seston carbon per day. These data indicate that
sphaeriid communities crop a significant portion of the
primary productivity of their small lentic habitats.
With extremely high filtering rates and dense populations (McMahon, 1999), C. fluminea is potentially a
major consumer of phytoplankton productivity. In the
Potomac River, phytoplankton densities and chlorophyll a concentrations declined as it flowed over dense
beds of C. fluminea, both measures falling 20 – 75%
lower than upstream values; current phytoplankton levels in this river section are considerably lower than
those prior to C. fluminea invasion. It was estimated
that the C. fluminea population filtered the entire water
column in the 3 – 4 day period required for it to pass the
river reach where the population was most dense (Cohen et al., 1984). Similar reduction in phytoplankton
density of a lotic habitat by C. fluminea has been reported by Leff et al. (1990). Reductions in phytoplankton density and chlorophyll a concentration must have
been due to clam filter feeding, particularly as discharge
volume variation, zooplankton feeding, toxic substances, and nutrient limitations were not different from
other river sections (Cohen et al. 1984). Similarly, Lauritsen (1986b) estimated that a C. fluminea population
of 350 clams / m2 in the Chowan River, NC, at an average depth of 5.25 m and clam filtering rate of
564 – 1010 mL / h, filtered the equivalent of the overlying water column every 1 – 1.6 days. At an average
depth of 0.25 m, current flow of 18.5 m / min, and
average clam filtering rate of 750 mL / h (Lauritsen,
1986b), the entire water column of the Clear Fork of
the Trinity River flowing over a C. fluminea population
with an average adult density of 3750 clams / m2 was filtered every 304 m of river reach, or every 16 min
(McMahon, unpublished observations). Such massive
filtering of the water column by dense bivalve populations could keep seston concentrations at low levels and
limit the energy available to other seston-feeding
species. Consumption of the majority of primary productivity by dense bivalve populations and the accumulation of that production in large, relatively long-lived,
predator-resistant adult clams may not only limit the
inorganic nutrients available for primary productivity
and the energy available to other primary consumers,
but may also divert energy flow from higher trophic
levels.
Bivalves, by filtering suspended seston, are water
clarifiers and organic-nutrient sinks. When co-cultured
with channel catfish, C. fluminea increased ambient
water O2 concentrations by reducing seston and turbidity levels (Buttner, 1986). Dense bivalve populations,
by removing phytoplankton and other suspended material, increase water clarity, and, by binding suspended
sediments into pseudofeces, accelerate sediment deposition rates (Prokopovich, 1969). Dense D. polymorpha
populations clarify water which increases light penetration, stimulating the growth of rooted aquatic macrophytes (Mackie and Schloesser, 1996).
Such major ecological impacts by D. polymorpha
have been observed in several locations in the lower
Great Lakes, St. Lawrence River and Hudson River
harboring dense mussel populations (reviewed by
MacIsaac, 1996). In these habitats, D. polymorpha
filtering increases water clarity. In the Hudson River,
D. polymorpha filtered the equivalent of the entire water column every 1.2 – 3.6 days (Strayer et al., 1999).
Phytoplankton densities are most negatively impacted
by D. polymorpha filtering in waters 1.85 m above
mussel beds (MacIsaac et al., 1999). Besides phytoplankton and small zooplankton, D. polymorpha filters
suspended clay and silt particles and binds them into
pseudofeces deposited in sediments (MacIsaac, 1996;
Strayer et al., 1999). Phytoplankton reductions by D.
polymorpha have been extensive, ranging from 59 to
91% in portions of the Great Lakes (MacIsaac, 1996)
and from 80 to 90% in the lower Hudson River, with
mussels less efficiently removing diatoms than other
smaller algae (Strayer et al., 1999). Filter feeding by D.
polymorpha can reduce bacterioplankton densities, but
can also stimulate bacterioplankton growth through
nutrient or organic carbon excretion (Cotner et. al.,
1995). Feeding by dense D. polymorpha populations
appears to favor development of cyanobacteria blooms
in some NA waters, perhaps due to reduction in N:P
ratio, enhanced light penetration, greater buoyancy of
cyanobacteria filaments, and / or chemical or mechanical inhibition of mussel filtering by cyanobacteria
(MacIsaac et al., 1996).
Zooplankton populations are reduced by D. polymorpha filtering. Small zooplankton such as rotifers
and even Dreissena veligers can be directly filtered;
rotifer density in western Lake Erie declining by 74%
within 5 years of D. polymorpha colonization, while
larger zooplankton species that avoid entrainment on
the mussel’s inhalant currents were relatively unaffected (MacIsaac, 1996). Similarly, zebra mussel
11. Mollusca: Bivalvia
invasion brought about 80 – 90% reductions in rotifers,
tintinnids, and copepod naupli densities in the lower
Hudson River (Strayer et al., 1999). Suppression of
small zooplankton could reduce fish populations whose
larvae utilize them as food and result in density increases of bacterioplankton on which small zooplankton feed (Strayer et al., 1999). Increased water clarity
and sediment nutrient levels appear to stimulate increased aquatic macrophyte growth in many areas colonized by D. polymorpha, favoring fish species associated with them over open-water species (MacIsaac,
1996). Increased macrophyte growth also appeared to
favor associated invertebrate species, whose densities
increased dramatically in the Hudson River after D.
polymorpha colonization while deep-water benthic invertebrates concurrently declined (Strayer et al., 1999).
Thus, overall, D. polymorpha populations appear to
divert resources from the pelagic zone and deep-water
sediments to vegetated shallows and zebra mussel beds
(Strayer et al., 1999), leading to increases in benthic invertebrate abundance and diversity due to habitat
structure enhancement among accumulated mussel
shells and increased food supply, enhancement of physical habitat appearing to have the greatest impact
(Botts et al., 1996). Increased bentic invertebrate densities support density increases in fish populations feeding on them (MacIsaac, 1996; Ricciardi et al., 1997;
Stewart et al., 1998; Strayer et al., 1999), These postcolonization impacts of D. polymorpha are extensive,
with almost all of 11 measured ecological variables being shifted by 50% after establishment of mussel
populations in the lower Hudson River (Strayer et al.,
1999). Increased food availability, associated with feeding on D. polymorpha, may also be responsible for
postzebra mussel colonization increases in diving-duck
flock sizes and staging durations in several portions of
Lake Erie (MacIsaac, 1996).
As freshwater clams filter suspended organic detritus and bacteria and consume sediment interstitial bacteria and organic detritus (Section III.D.2), they may be
significant members of the aquatic decomposer assemblage. Pisidium casertanum and P. corneum feed primarily by filtering sediment bacteria (Lopez and
Holopainen, 1987). Hornbach et al. (1982, 1984b) estimated that only 24 – 35% of energy needs of a population of Sphaerium striatinum were met by filter feeding,
the remaining coming from sediment organic detritus.
C. fluminea pedally feeds on sediment detritus (Reid et
al., 1992), impacting the decomposer community (Hakenkamp and Palmer, 1999). Musculium transversum
utilizes its long inhalant siphon to vacuum organic detritus from the sediment surface (Way, 1989). The detritivorous habit of many freshwater bivalve species may
divert primary productivity ordinarily lost to respiration
389
of the detritivorous community back into bivalve tissue
where it is made re-available to higher trophic levels.
The activity of freshwater bivalves may also directly affect habitat physical characteristics. Deposition
of calcium in shells may reduce ambient Ca concentrations. In a C. fluminea population of 32 clams / m2, annual fixation of shell CaCO3 was 0.32 kg CaCO3 /m2
per year (Aldridge and McMahon, 1978). At this rate,
shell CaCO3 fixation in a dense population of 100,000
clams / m2 (Eng, 1979) could be 50 – 60 kg Ca CO3 /m2
per year. Such rapid removal of calcium to shells accumulating in the sediments could reduce water hardness,
particularly in lentic habitats (Huebner et al., 1990).
Seasonal cycles of shell growth could induce seasonal
cycles in water Ca concentration, being greatest in winter when shell growth is minimal and least in summer
when shell growth is maximal (Rooke and Mackie,
1984c). Bivalves can also affect solute flux between
sediments and the water column. Unionoidean mussels
enhanced release of nitrate and chloride and inhibited
CaCO3 release from sediments (Matisoff et al., 1985).
Dense sphaeriid populations can be the principal affectors of sediment dissolved O2 demand, in cases reaching levels characteristic of semipolluted and polluted
streams (Butts and Sparks, 1982). D. polymorpha and
C. fluminea increase sedimentation rates, negatively or
positively impact phosphate and nitrate fluxes between
sediments and the water column, change the N:P ratio
of the water column, clarify water, and change sediment physical make up by accumulation of their shells
(Arnott and Vanni, 1996; Strayer et al., 1999).
Small bivalves (sphaeriids, juvenile unionoideans,
D. polymorpha, and C. fluminea) can be a major food
source for third trophic level carnivores, including fish
and crayfish (Section III.C.3). Bivalve flesh has a low
caloric content, 3.53 – 5.76 kcal / g ash-free dry weight
(Wissing et al., 1982), reflecting low lipid:protein ratios. C. fluminea has tissue organic C:N ratios ranging
from 4.9 – 6.1:1, indicative of a 51 – 63% of dry weight
protein content (Williams and McMahon, 1989). High
protein content makes bivalve flesh an excellent food
source to sustain predator tissue growth.
Many freshwater bivalve populations are highly
productive (particularly sphaerids, C. fluminea, and D.
polymorpha), rapidly converting primary productivity
into tissue energy available to third trophic level carnivores. Productivity values range from 0.019 g C / m2
per year for a Pisidium crassum population to 10.3 g
C / m2 per year for a C. fluminea population (Aldridge
and McMahon, 1978) and 75 g C / m2 per year for a
D. polymorpha population (Mackie and Schloesser,
1996); the average for 11 sphaeriid species and two
species of Corbicula was 2.5 g C / m2 per year (Burky,
1983). Other values include 12.8 – 14.6 g C / m2 per
390
Robert F. McMahon and Arthur E. Bogan
year for an Arizona C. fluminea canal population
(computed from data of Marsh, 1985) and 1.07 g
C / m2 per year for a New Brunswick lake unionoidean
population of Elliptio complanata (computed from
data of Paterson, 1985).
Bivalves are generally more efficient at converting
assimilated food energy into new tissue than most
second trophic level aquatic animals because their sessile, filter-feeding habits minimize energy expended in
food acquisition, allowing efficient transfer of energy
from primary production to bivalve predators. Thus,
bivalves can be important conduits of energy fixed by
phytoplankton photosynthesis to higher trophic levels.
However, such trophic energy transfer is mainly
through smaller species and juvenile specimens, as large
adults are relatively immune to predation (Section 3.C).
The measure of efficiency of conversion of assimilated energy absorbed across the gut wall into energy
fixed in new tissue growth is net growth efficiency:
% Net Growth Efficiency
P
A(100)
where P is the productivity rate or rate of energy or organic carbon fixation into new tissue growth by an individual or population and A the assimilation rate or
rate of energy or organic carbon assimilated by an individual or population. The greater the net growth efficiency of a second trophic level species, the more efficient its conversion of assimilated energy into flesh,
and, therefore, the greater the potential for energy flow
though it to third trophic level predators (reviewed by
Russell-Hunter and Buckley, 1983; Holopainen and
Hanski, 1986; Hornbach, 1985). Net growth efficiencies (Table III), are 9 – 79% in sphaeriids (average
55%) and 58 – 89% in C. fluminea, indicating efficient
conversion of assimilated energy into flesh compared to
other aquatic second trophic level animals.
5. Bivalves as Biomonitors
Freshwater bivalves, particularly unionoideans C.
fluminea and D. polymorpha, have characteristics such
as long life spans, and growth and reproductive rates
sensitive to environmental perturbation (Burky, 1983)
that make them good biomonitors (Imlay, 1982). They
can be held in field enclosures without excessive maintenance. Shell growth is sensitive to environmental
variation and / or disturbance (Way and Wissing, 1982;
Fritz and Lutz, 1986; Belanger et al., 1986a, b; Way,
1988), and the valves remain as evidence of death, allowing mortality rates to be estimated. Shells can be
marked by tags (Rosenthal, 1969; Young and Williams,
1983c), including floating tags tethered to the shell
(Englund and Heino, 1994b), paint, or shell-etching
(McMahon and Williams, 1986a).
Bivalves can be collected throughout the year and
easily shipped alive long distances. Because large
species stay in place (exception is C. fluminea), they experience conditions in the monitored environment
throughout life (Imlay, 1982). Annual shell-growth increments allows determination of variation in heavymetal pollutant levels over long periods by analysis of
levels in successively secreted shell layers (Imlay, 1982).
However, there is controversy regarding use of shellgrowth rings to age unionoideans. Growth rings are
produced annually in some species (Neves and Moyer,
1988), but not in others (Downing et al., 1992). Large
size allows analysis of pollutant levels in single individuals, and the wide geographical ranges of some species
(Imlay, 1982) allow comparisons across drainage systems. Table VI indicates the types of pollutants and environmental perturbations monitored by freshwater bivalves. The broad distributions of C. fluminea and D.
polymorpha in North America and their dense, readily
collectible populations make them excellent biomonitors compared to unionoideans, many species of which
have restricted distributions and / or are threatened or
endangered (Section III.A).
D. Evolutionary Relationships
Sphaeriid, dreissenids, corbiculids, and unionoideans represent separate evolutionary invasions of
freshwaters. Sphaeriids, dreissenids, and corbiculids fall
in the order Veneroida, and unionoideans in the order
Unionoida (Allen, 1985). The Veneroida became successful infaunal filter feeders though evolution of inhalant and exhalant siphons, allowing maintenance of
respiratory and feeding currents while burrowed. Their
eulamellibranch gills with fused filaments (Fig. 5) were
a clear advancement over the primitive (i.e., filibranch)
condition with separate gill filaments attached by interlocking cilia (Allen, 1985).
The family, Sphaeriidae, in the superfamily, Corbiculacea, with a long freshwater fossil history extending
from the Cretaceous (Keen and Dance, 1969) has
evolved along two major lines. The first, represented by
Pisidium, involves adaptation to life in organically rich
sediments by filtering interstitial bacteria (Lopez and
Holopainen, 1987), making them dominant in profundal habitats. The second, represented by Sphaerium
and Musculium, involves adaptation to life in small,
shallow, lentic or lotic habitats subject to predictable
seasonal perturbation, such as habitat-drying. Their
adaptations include estivation during prolonged emergence (Section III.A). Like many of the Veneroida, the
byssus is absent in adults of most species.
The superfamily Unionoidea, like the Sphaeriidae,
has a long freshwater fossil history, extending from at
11. Mollusca: Bivalvia
391
TABLE VI
List of some Investigations Involving Utilization of Bivalves to Monitor Effects or Levels of Pollutants in
Freshwater Habitats
Pollutant monitored
Species utilized
Literature citations
Arsenic
Corbicula fluminea
Cadmium
Eight unionoidean species,
Anodonta anatina
Anodonta cygnea
Corbicula fluminea
Chromium
Copper
Eight unionoidean species
Corbicula fluminea
Corbicula fluminea
Iron
Lead
Lamellidens marginalis
Corbicula fluminea
Corbicula fluminea
Elder and Mattraw (1984), Price and
Knight (1978), Tatem, (1986)
Price and Knight (1978)
Hemelraad et al. (1985)
Hemelraad et al. (1985)
Elder and Mattraw (1984), Graney et al. (1984),
Price and Knight (1978), Tatem (1986)
Price and Knight (1978)
Elder and Mattraw (1984), Tatem (1986)
Annis and Belanger (1986), Elder and Mattraw (1984),
Price and Knight (1978), Tatem (1986)
Hameed and Raj (1990)
Tatem (1986)
Annis and Belanger (1986), Elder and Mattraw (1984),
Price and Knight (1978), Tatem (1986)
Hameed and Raj (1990)
Price and Knight (1978)
Elder and Mattraw (1984), Tatem (1986)
Hameed and Raj (1990)
Elder and Mattraw (1984), Price and Knight (1978)
Hameed and Raj (1990)
Price and Knight (1978)
Herwig et al. (1985)
Belanger et al. (1986b), Elder and Mattraw (1984),
Foe and Knight (1986c)
Hameed and Raj (1990)
Belanger et al. (1986a, 1987)
Elder and Mattraw (1984), Tatem (1986)
Elder and Mattraw (1984), Tatem (1986)
Pugsely et al. (1985)
Fisher et al.1993
Borcherding (1992), Fisher et al. (1993)
Elder and Mattraw (1984), Hartley and
Johnston (1983), Tatem (1986)
Foe and Knight (1986c), Horne and
MacIntosh (1979), Weber (1973)
Dreier and Tranquilli (1981), Ferris et al. (1988),
Foe and Knight (1987), McMahon and Williams (1986b)
Borcherding and Volpers (1994)
Englund and Heino (1994a)
Englund and Heino (1994a)
Manganese
Mercury
Nickel
Tin
Zinc
Lamellidens marginalis
Eight unionoidean species
Corbicula fluminea
Lamellidens marginalis
Corbicula fluminea
Lamellidens marginalis
Eight unionoidean species
Anodonta sp.
Corbicula fluminea
Pentachlorophenol (PCP)
Pesticides
Lamellidens marginalis
Corbicula fluminea
Lampsilis siliquiodea
Corbicula fluminea
Lampsilis siliquoidea
Dreissena polymorpha
Dreissena. Polymorpha
Corbicula fluminea
Sewage effluents
Corbicula fluminea
Power station effluents
Corbicula fluminea
Natural waters
Dreissena polymorpha
Unio tumidus
Anodonta anatina
Asbestos
Octachlorostyrene
Polychlorinated
Biphenols (PCBs)
least the Triassic (Haas, 1969). A long fossil history and
tendency for reproductive isolation due to gonochorism
and the parasitic glochidial stage (Section III.A) which
allows sympatric speciation via new glochidial fish host
acquisition (Graf, 1997) has led to extensive radiation,
the Unionoidea being represented worldwide by 150
genera and a great number of species (Allen, 1985).
Their origin remains unclear. Similarity of shell structure suggests a relationship to marine fossil species in
the order Trigonoida (Allen, 1985). Like the majority of
sphaeriids, adult unionoideans lack a byssus, which
would be of no use in their infaunal habitats. They
dominant the shallow regions of larger, relatively
stable lentic and lotic habitats, particularly in stable
sand – gravel sediments (Section III.A). The division of
the superfamily, Unionoidea, into the presumed more
primitive family, Margaritiferidae, and the more derived
family, Unionidae, has been partially supported by mitochondrial DNA analysis of NA species. This study
also supported the monophyly of the NA unionoidean
subfamilies Anodontinae and Ambleminae and tribe
Pleuorblemini, but provided only weak support for the
392
Robert F. McMahon and Arthur E. Bogan
tribe Lampsilini (Lydeard et al., 1996). Further molecular studies are needed to elucidate the complex phylogeny of the Unionoidea, including analysis of species
outside of North America.
The Sphaeriidae and Unionoidea, with long freshwater fossil histories, have evolved distinct niches.
While both occur in stable habitats, the majority of
sphaeriids inhabit smaller ponds and streams or the
profundal regions of lakes, and the majority of
unionoideans inhabit stable sediments of shallow portions of larger rivers and lakes (Sections III.A and III.B).
C. fluminea entered freshwater only in recent (Pleistocene) times (Keen and Casey, 1969), making it unlikely to complete effectively with sphaeriids or
unionoideans whose long fossil histories have allowed
them to become highly adapted to their preferred stable
freshwater habitats. Indeed, C. fluminea does not appear to seriously impact native NA bivalves in undisturbed freshwaters (McMahon, 1999; Section C.3).
Rather, it is adapted for life in unstable habitats — subject to periodic catastrophic perturbation, particularly
flood-induced sediment disturbance — which are generally unsuitable for sphaeriids or unionoideans (Section
III.A). Among its adaptations to high-flow habitats are:
(1) capacity for rapid burrowing; (2) a strong, heavy,
concentrically sculptured, inflated shell; and (3) juvenile
retention of a byssal thread — all of which allow maintenance of position in sediments (Vermeij and Dudley,
1985). Additionally, its high fecundity, elevated growth
rate, early maturity, and extensive capacity for dispersal
(Section III.B) allow it to colonize habitats from which
other bivalves have been extirpated and rapidly recover
from catastrophic population reductions. Thus, C. fluminea evolved a niche in unstable, disturbed habitats
not utilized by sphaeriids or unionoideans, having
evolved from an estuarine Corbicula ancestor inhabiting similar environments in the upper, near-freshwater
portions of estuaries (Morton, 1982).
The superfamily Dreissenacea, containing D. polymorpha, while superficially resembling marine mytilacean mussels (order Mytiloida), is placed in the Veneroida because of its advanced eulamellibranch
ctenidium. Adult dreissenaceans retain an attachment
byssus, considered a primitive condition. In contrast,
the mytiliform shell with its reduction of the anterior
valves and anterior adductor muscle, is a derived adaptation to an epibenthic niche characterized by byssal attachment to hard surfaces (Allen, 1985; Mackie and
Schloesser, 1996). D. polymorpha probably evolved
from an estuarine dreissenacean ancestor of the genus
Mytilopsis. As with C. fluminea, its fossil record indicates a recent, Pleistocene introduction to freshwaters
(Mackie et al., 1989). Also, like C. fluminea, D. polymorpha appears to have successfully colonized freshwaters because its niche (i.e., an epibenthic species
attached to hard substrates) is not occupied by infaunal,
burrowing sphaeriids or unionoideans, thus minimizing
competition with these more advanced groups.
Freshwater bivalves have characteristics that are
uncommon in comparable marine species. Most shallow-burrowing marine species are gonochoristic with
external fertilization and free-swimming planktonic
veligers. A planktonic veliger is nonadaptive in lotic
freshwater habitats, as it would be carried downstream
before settlement, eliminating upstream populations.
Thus, the planktonic veliger stage is suppressed in
sphaeriids, which brood embryos in gill marsupia and
release large, fully formed juveniles ready to take up life
in the sediment. In unionids, eggs are retained in gill
marsupia and hatch into the glochidium, whose parasitism of fish hosts allows upstream as well as downstream dispersal and permits release of well-developed
juveniles into favorable habitats (Section III.B). Even the
recently evolved C. fluminea has suppressed the planktonic veliger stage of its immediate estuarine ancestors
(Morton, 1982), producing a small, but fully formed juvenile whose byssal thread allows immediate settlement
and attachment to the subtrate. Retention of external
fertilization and a free-swimming veliger in D. polymorpha are primitive characteristics reflecting its recent
evolution from an estuarine ancestor.
Downstream veliger dispersal would preclude D.
polymorpha from establishing populations in high-flow
lotic habitats, unless upstream impoundments provide
replacement stock. In fact, in Europe and Asia, this
species is most successful in large, lentic or low-flow
lotic habitats such as canals or large rivers (Mackie et
al., 1989). Serial impoundment of rivers has provided
lentic refugia for this species in otherwise lotic
drainages, facilitating its invasion of NA waters
(Mackie and Schloesser, 1996).
Another common adaptation in freshwater species is
hermapharoditism with a capacity for self-fertilization.
Hermaphroditism makes all individuals in a population
reproductive, allowing rapid population expansion
during favorable conditions. Hermaphroditism allows
sphaeriids to re-establish populations after predictable
seasonal perturbation and C. fluminea, along with its
high fecundity, to rapidly re-establish populations after
catastrophic habitat disturbance. It makes both sphaeriids and C. fluminea invasive, as the introduction of a
single individual can found a new population. In contrast, the gonochorism of most unionoideans and D.
polymorpha limits their invasive capacity, reflected by
their stable, relatively undisturbed habitats. Hermaphroditism occurs in only seven species within three subfamilies of NA Unionoidea (Hoeh et al., 1995) with widely
different habitats and reproductive traits. Selection pressures for evolution of hermaphroditism within these
species have not been elucidated.
11. Mollusca: Bivalvia
Finally, unionoideans and sphaeriids have different
shell morphologies relative to comparable shallowburrowing marine bivalves. Freshwater species generally do not have denticulated or crenulated inner-valve
margins, extensive radial or concentric shell ridges,
tightly sealing valve margins, well-developed hinge
teeth, overlapping shell margins, or uniformally thick,
inflated, strong shells to the degree displayed by shallow-burrowing marine species (Vermeij and Dudley,
1985). These shell characteristics allow shallowburrowing marine species to maintain position in unstable substrates and / or resist shell cracking or boring
by large predators common in marine habitats. Lack of
such structures in unionoideans and sphaeriids reflects
their preference for stable sediments and the general
absence of effective predators on adult freshwater bivalves; certainly there are no shell-boring predators of
NA freshwater species. Indeed, the shells of freshwater
unionoideans display considerably less nonlethal,
predator-induced damage than shallow-burrowing marine species (Vermeij and Dudley, 1985). While retention by C. fluminea of a thick, extremely strong shell
(Kennedy and Blundon, 1983; Miller et al., 1992),
lacking pedal and siphonal gapes, with a well-developed hinge, shell inflation, and concentric ornamentation, may represent a primitive condition, it allows this
species to inhabit sediments too unstable to support
unionoideans or sphaeriids. Such primitive characteristics reflect similar adaptations in the immediate estuarine ancestor of this recently evolved freshwater species.
In contrast, the thin, fragile shell of D. polymorpha
(Miller et al., 1992) may have evolved due to byssal attachment of the species to hard substrates where the
shell is not subject to damage from unstable sediments.
IV. COLLECTING, PREPARATION FOR
IDENTIFICATION, AND REARING
A. Collecting
Small sphaeriid clams are best collected by removing
sediments containing clams with: (1) a trowel or shovel
(shallow water); and (2) a long-handled dip net or shell
scoop with a mesh of 0.35 mm (moderate depths) or
by Ekman, Ponar, or Peterson dredges (deeper water),
the heavier Ponar and Peterson dredges being better in
lotic habitats. Drag dredges must have sediment catch
bags of small mesh (1 mm or less). Use of scuba is also
an effective means of collecting sphaeriids. Larger
sphaeriids species can be separated from fine sediments
with a 1-mm mesh sieve, but a 0.35-mesh is required for
smaller specimens. Fragile sphaeriids should be separated from sediments by gentle vertical agitation of the
sieve at the water surface to prevent shell damage.
393
Coarse material should be removed prior to sieve
agitation to avoid shell breakage. Exceptionally small,
fragile species can be collected by washing small quantities of sediment into settlement pans, specimens being
revealed after sediments settle and the water clears.
Ekman, Ponar, or Peterson dredges are best for
quantitative samples of sphaeriids, because they remove sediments from under specific sediment surface
areas. Quadrat frames can be placed on the bottom
(Miller and Payne, 1988) and all surface sediments
within the frame collected for sorting. Such frames can
be utilized in shallow or deeper waters (the latter in
conjunction with scuba equipment). Drag dredging at
specified speeds for known intervals allows partial sample quantification. Core sampling devices consisting of
tubes driven into the substrate to specific depths can
yield both accurate estimates of sphaeriid density and
sediment depth distributions. Core samplers sample
only a small surface area, thus are best utilized with
dense populations and / or repetitive sampling.
Low densities of some unionoidean populations
make their collection difficult. Where populations are
dense, shoveling sediments through a sieve allows collection of a broad range of sizes and age classes (Miller
and Payne, 1988). As unionoideans are larger than
sphaeriids, sieve mesh sizes of 0.5 – 1 cm may be appropriate unless recently settled juveniles must be collected.
In less dense, shallow-water populations, hand-picking
using a glass-bottomed bucket to locate specimens can
often suffice. In turbid waters and / or soft sediments
(sand or mud), shell rakes may be utilized. Hand-picking and shell rakes can select larger, older individuals. In
shallow, turbid water, using hands and fingers to systematically feel for mussels buried in soft sediments can
be effective, but selects larger individuals and can lead
to cut fingers. On rocky or boulder bottoms,
unionoideans generally accumulate in crevices or near
downstream bases of large rocks. Here, only hand-picking (in conjunction with scuba techniques in deeper water) is effective. Unionoideans may also be collected by
drag or brail dredges. Mussel brails (often used by commercial shellers) consist of a bar with attached lines terminating in blunt-tipped gang hooks. When dragged
over unionid beds, mussels clamp the valves onto the
hooks allowing them to be brought to the surface.
Quantitative sampling of unionoideans can be accomplished with quadrats, along with scuba in deeper
waters (Isom and Gooch, 1986; Miller and Payne,
1988; Waller et al., 1993). Only heavy Ponar and Peterson dredges bite deeply enough into the substrate to
take large unionoidean species. In sparse populations,
errors in density estimates can result from the small
surface areas that these dredges sample, requiring
numerous samples to improve accuracy. Drag dredges
sample larger areas and may be partially quantitative,
394
Robert F. McMahon and Arthur E. Bogan
but generally do not provide accurate estimates. A
skimmer dredge collected 62.3% of mussels in its path,
but resulted in 10% mortality of thin-shelled species
(Miller et al., 1989) Unionoideans may also be
collected during natural or planned water level drawdowns (White, 1979) by collecting either emersed individuals or those migrating downshore and accumulating at the water’s edge.
In a comparative study of sampling, quadrat sampling provided the most accurate analysis of a
unionoidean community, but was difficult and expensive to carry out in deeper waters, where systematic
sampling by scuba produced the best results (Isom and
Gooch, 1986). In a comparison of the accuracy of
quadrat and qualitative timed searches for unionoideans, it was found that timed searches overestimated
large species, highly sculptured species and those whose
shells protruded from the substrate, while underestimating buried, small, smooth-shelled species. In contrast, quadrat sampling tended to underestimate rare
species and total number of species unless a large number of quadrats were sampled (Vaughn et al., 1997).
Extensive sampling may be required to find rare
species, particularly if their population densities need
to be assessed; thus, time and funding limitations may
prevent accurate assessment of presence / absence and
population densities of rare unionoideans (Kovalak et
al., 1986; Vaughn et al., 1997). Accurate sampling of
rare unionoidean species is required for their effective
management and conservation. As many NA unionoidean species are endangered, one should ascertain the
status of any species before permanently removing
specimens from their natural habitats. Take only dead
shells or, if absolutely necessary, a few living specimens
for identification.
C. fluminea is easily collected because it occurs in
high densities, prefers shallow waters, and is easily
identified. Individuals can be separated from sediments
with a 1-mm mesh sieve. Qualitative samples are best
obtained by shovel or drag dredge (Williams and
McMahon, 1986) and quantitative samples by quadrat
frame (Miller and Payne, 1988), or Peterson (Aldridge
and McMahon, 1978) or Ekman dredges (Williams
and McMahon, 1986). In rock or gravel substrates, C.
fluminea are best taken by hand, hand trowel, or
shovel; collected sediments should be passed though a
coarse mesh sieve to separate specimens. In fast-flowing
streams, specimens of C. fluminea accumulate in
crevices or behind the downstream sides of large rocks.
D. polymorpha is best collected by scuba with
manual removal, using quadrat frames for quantification, because of its byssal attachment to hard surfaces
and its preference for deeper waters (1 – 2 m).
Ripping of individuals from the byssus damages their
tissues, thus the byssus should be cut with a knife or
sharp trowel before removal. Juveniles can be collected
after they have settled on settlement blocks or submerged buoys set out during the reproductive season.
Ekman, Ponar, Peterson, and drag dredges are generally
not suitable for collection of attached epibenthic
species such as D. polymorpha, but they can be used
for populations on soft or semisoft benthic substrates
(Hunter and Baily, 1992).
Juveniles of sphaeriids and C. fluminea can be surgically removed from brood sacs or collected from sediments after release with a fine mesh sieve (mesh size
0.35 mm). Juvenile C. fluminea can also be obtained
by release from freshly collected, gravid adults left in
water for 12 – 24 h (Aldridge and McMahon, 1978).
Planktonic C. fluminea juveniles and D. polymorpha
veligers may be taken with a zooplankton net towed
behind a boat or held in current flow. Their densities
may be assessed by passing known volumes of water
through a zooplankton net. Plankton net mesh size for
collection of C. fluminea juveniles should be 200 m
⬃40 m for all planktonic stages of D. polymorand
pha, including the trochophore. Recently settled juveniles of unionoideans may be sieved from sediments.
Glochidia can be surgically removed from demibranchs
of gravid females or encysted glochicia taken from the
fins, pharyngeal cavity, or gills of their fish hosts (for a
list of unionoidean fish hosts, see Watters, 1994b).
B. Preparation for Identification
To preserve bivalves, larger individuals should first
be narcotized or relaxed, allowing tissues to be preserved in a lifelike state and preservatives to penetrate
tissues through gaped shell valves. Live bivalves placed
directly in fixatives clamp the valves, which prevents
preservative penetration. There are a number of bivalve
relaxing agents (Coney, 1993; Araujo et al., 1995), including alcoholized water (either 3% ethyl alcohol by
volume with water or 70% ethyl alcohol added slowly,
drop by drop, to the medium until bivalves gape), chloroform added slowly to the medium, methol crystals
(one level teaspoon per liter, scattered on the water surface), propylene phenoxetol and phenoxetol BPC
(5 mL of product emulsed with 15 – 20 mL of water,
added to water containing bivalves or introduction of a
droplet equal to 1% of holding water volume), phenobarbitol added in small amounts to the holding
medium, magnesium sulfate (introduced into holding
medium over a period of several hours to form a
20 – 30% solution by weight), magnesium chloride
(7.5% solution by weight), and urethane.
None of these agents works equally well with all
species. Laboratory tests of narcotizing agents against
11. Mollusca: Bivalvia
C. fluminea revealed propylene phenoxetol to be the
only agent capable of relaxing this species for experimental surgery, and allowing recovery on return to
fresh medium (Kropf-Gomez and McMahon, unpublished). Heating bivalves to 50°C for 30 – 60 min causes
most species to relax and gape widely, but is lethal. In
larger specimens, wooden pegs or portions of matchsticks forced between the valves prior to fixation allows
preservative penetration of tissues.
The best long-term preservative for freshwater bivalves is 70% ethyl alcohol (by volume with water).
Specimens may be initially fixed in 5 – 10% formaldehyde solutions (by volume with 40% formaldehyde solutions) for 3 – 7 days. But formaldehyde is acidic and
dissolves the calcareous portion of shells unless pHneutralized by the addition of powdered calcium carbonate (CaCO3) to make a saturated solution, 5 g of
powdered sodium bicarbonate (NaHCO3) per liter, or
1.65 g of potassium dihydrogen orthophosphate and
7.75 g disodium hydrogen orthophosphate per liter
(Smith and Kershaw, 1979). After 3 – 7 days in
formaldehyde, specimens should be transferred to 70%
ethyl alcohol for permanent preservation. Addition of
1 – 3% glycerin (by volume) to alcohol preservatives
keeps tissues soft and pliable (Smith and Kershaw,
1979). Smaller species (shell length 15 mm) generally
do not require relaxation. Tissues to be utilized in microscopy should be preserved in gluteraldehyde or
Bouin’s solution.
Shells can be cleaned with a mild soap solution and
soft brush. Organic material can be digested from shell
surfaces by immersion in a dilute (3% by weight with
water) solution of sodium or potassium hydroxide at
70 – 80°C, thereafter, removing remaining organic
matter with a soft brush. For dry-keeping, the shell periostracal surface should be varnished or covered with
petroleum jelly to prevent drying, cracking, and / or
peeling. Numbers identifying collection and specimen
can be marked on the inner shell surface with India ink.
In order to remove soft parts from living bivalves,
immerse them in water at 60°C and remove tissues
after valves fully gape. Separated flesh can be fixed in
70% alcohol. For fragile sphaeriids, flesh is best removed with the tip of a fine needle, manipulating specimens with a fine brush. Shells should be dried in air at
room temperature, not in an oven; heat causes shells to
crack and their periostracum to crack and peel.
Both soft tissue and shell characteristics are utilized
in the identification of freshwater bivalves, so both
must be preserved for species identification. For
unionoideans, C. fluminea, and D. polymorpha, most
diagnostic taxonomic characteristics can be seen by eye
or with a 10 hand lens. For sphaeriids, a dissecting
microscope with at least 10 – 30 power or a com-
395
pound microscope is required (Ellis, 1978). Anatomical
details are best observed on dry shells or soft tissues
immersed in water.
Identification of recently released juvenile bivalves
is difficult and may require preparation of stained slide
whole mounts. Glochidia are best identified by removal
from a gravid adult of an identified species as are juvenile sphaeriids. Only the juveniles of C. fluminea and
larval stages of D. polymorpha are routinely found in
the plankton. The juvenile of C. fluminea (Fig. 15A) is
easily recognizable, and can be readily separated from
the pediveliger of D. polymorpha by the presence of a
fully formed foot, lack of a velum and a D-shaped shell
while the foot is formed only in the pediveliger stage of
D. polymorpha which has an umbonal shell (Nichols
and Black, 1994). The planktonic veliger of D. polymorpha is clearly distinguishable by its ciliated velum,
lack of a foot and D-shaped shell (Nichols and Black,
1994; Fig. 15C). The glochidia of many unionoidean
species have specific fish species hosts (Watters, 1994b),
whose identification will assist glochidial identification.
Identification of glochidia without knowledge of the
host fish or species of origin is extremely difficult, characters such as presence / absence of attachment hooks
or threads and glochidial size may allow assignment to
a family or subfamily, but may be unreliable even at
these higher taxonomic levels (Bauer, 1994).
C. Rearing Freshwater Bivalves
For artificial rearing of freshwater bivalves, water in
holding tanks should be temperature-regulated. Adequate aeration, filtration, and ammonia removal systems
are required, as some species have low hypoxia and ammonia tolerances (Byrne et al. 1991a, 1991b).
As unionoideans and C. fluminea are filter phytoplankton, they require a constant supply of filterable
food to remain healthy and growing. The best artificial
food appears to be algal cultures. When fed monoalgal cultures of the green algae, Ankistrodesmus and
Chlorella vulgaris or the cyanobacteruim, Anabaena
oscillariodes, assimilation efficiencies in C. fluminea
were 47 – 57% and net growth efficiencies, 59 – 78%,
making them excellent food sources for this species
(Lauritsen, 1986a).
Artificial diets do not appear to be as successful as
algal cultures in maintaining C. fluminea growth. When
fed either ground nine-grain cereal, rice flour, rye, bran,
brewers’ yeast, or artificial trout food, small C. fluminea
(5 – 8 mm SL) starved on all but nine-grain cereal, the
latter supporting little tissue growth. Supplementing
these grain diets with live green algae (Ankistrodesmus
sp.) greatly enhanced tissue growth, but the greatest
growth occurred on pure Ankistrodesmus cultures (Foe
396
Robert F. McMahon and Arthur E. Bogan
and Knight, 1986b). C. fluminea fed mixed cultures of
the green algae Pedinomonas sp., Ankistrodesmus sp.,
Chlamydomonas sp., Chorella sp., Scenedesmus sp., and
Selenastrum sp. had maximal growth when mixtures did
not include Selenastrum, which was toxic to this species.
Greatest tissue growth occurred in clams fed mixed cultures of all five remaining algal species. Growth declined
with number of algal species in feeding cultures; feeding
with two algal species resulting in starvation (Foe and
Knight, 1986b). Thus, artificial bivalve culture appears
to be best supported on mixed algal diets, but certain
toxic algal species must be avoided.
Temperature also affects bivalve growth rate.
When 5 – 8 mm SL specimens of C. fluminea were fed
mixed algal cultures of Chamydomonas, Chlorella, and
Ankistrodesmus at 105 cells / mL, assimilation efficiencies were maximal at 16 and 20°C (48 – 51%). Tissue
growth was maximal at 18 – 20°C and became negative
(tissue loss) at 30°C (Foe and Knight, 1986a), suggesting 18 – 20°C to be an ideal culture temperature for
this species and, perhaps, other NA bivalves. In algaefed cultures of D. polymorpha, growth was maximal at
15°C and suppressed at 20°C (Walz, 1978b). However,
tissue growth in a natural population of C. fluminea increased up to 30°C (McMahon and Williams, 1986a),
indicating that artificial culture systems are not equivalent to field conditions in supporting bivalve growth. In
this regard, excellent tissue growth in C. fluminea was
supported by algal cultures produced by several days’
exposure to sunlight of water taken from the natural
habitat of the clam to increase its algal concentration
(Foe and Knight, 1985). Thus, ideal culture conditions
for unionoideans and C. fluminea would appear to be a
20°C holding temperature and feeding with natural algal assemblages whose growth has been promoted with
inorganic nutrients and exposure to sunlight; maximal
growth being achieved in D. polymorpha under the
same conditions at 15°C.
Specimens of C. fluminea, D. polymorpha and
unionoideans may be held for long periods in the laboratory without feeding. C. fluminea survived 154 days
of starvation at room temperature (22 – 24°C) while
sustaining tissue weight losses ranging from 41 to 71%
(Cleland et al., 1986). Similarly, I have held unionoideans in the laboratory for many months without
feeding. Samples of D. polymorpha experienced 100%
mortality after being held in the laboratory without
feeding for 166, 514, and 945 days at 25, 15, and 5°C,
respectively (Chase-Off and McMahon, unpublished
data). Thus, maintenance at low temperature (10°C)
greatly prolongs the time bivalves may be held in good
condition without feeding.
C. fluminea has never been reared successfully to
maturity or carried through a reproductive cycle in artificial culture, although field-collected, nongravid
adults released juveniles after four months in laboratory culture (King et al., 1986). The glochidium stage
makes unionoideans difficult to rear in the laboratory
as it requires encystment in a fish host for successful
juvenile metamorphosis, but it can be accomplished
(Young and Williams, 1984a). Application of an immunosuppressant agent has allowed glochidial transformation to juveniles on nonhost fish where they
would otherwise be rejected by the immune system of
the host fish (Kirk and Layzer, 1997). Glochidia of several unionoidean species have been transformed into
juveniles in vitro in a culture medium containing physiologic salts, amino acids, glucose, vitamins, antibiotics,
and host fish plasma (Isom and Hudson, 1982). Juvenile utterbackia imbecillis and Epioblasma triquetra
have been successfully cultured in a medium of river
water exposed to sunlight for 1 – 4 days to enhance algal concentration. Addition of silt enhanced juvenile
growth in both species, while feeding artificial, mixed
cultures of three algal species resulted in starvation
(Hudson and Isom, 1984), an observation supported
by recent studies suggesting that juvenile unionoideans
feed primarily on interstitial water drawn from sediments through the pedal gape, such that silt and associated microdetritus and bacteria may be their main ingested food source (Yeager et al., 1994).
Many sphaeriid species can be easily maintained in
simple artificial culture systems. Ease of artificial culture
in this group may relate to their feeding on sediment
organic detritus (Burky et al., 1985b; Hornbach et al.,
1984b) and interstitial bacteria (Lopez and Holopainen,
1987), making use of algal cultures as a food source unnecessary. Hornbach and Childers (1987) maintained
Musculium partumeium through successful reproduction in beakers with 325 mL of filtered river water,
without sediments, on a diet of 0.1 mg of finely ground
Tetra Min© fish food/clam per day. The first generation
in this simple culture system survived 380 – 500 days, a
life span equivalent to the natural population (Hornbach et al., 1980). Similarly, Rooke and Mackie (1984c)
maintained 20 adult Pisidium casertanum in a 6-L
aquaria with sediments for 35 weeks without feeding,
suggesting that individuals fed on sediment bacteria or
organic deposits.
Live molluscs rapidly remove dissolved calcium
from culture media (Rooke and Mackie, 1984c), thus
calcium levels in holding media should be augmented
by the addition of CaCO3. The ease with which
sphaeriids can be artificially cultured makes them ideal
for laboratory microcosm experiments. Ideal culture
conditions appear to include provision of natural
sediments, a source of calcium (i.e., ground CaCO3),
and finely ground food of reasonable protein content,
such as aquarium fish food or brewers’ yeast, which
may be directly assimilated by clams or support the
11. Mollusca: Bivalvia
growth of interstitial bacteria upon which they feed
(Lopez and Holopainen, 1987).
V. IDENTIFICATION OF THE FRESHWATER
BIVALVES OF NORTH AMERICA
A. Taxonomic Key to the Superfamilies
of Freshwater Bivalvia
There are five bivalve superfamilies with freshwater
representatives in North America. Of these the
Unionoidea, Corbiculoidea, and Dreissenoidea contain
397
the true freshwater species and comprise the vast
majority of freshwater bivalve fauna. The remaining
two superfamilies, Cyrenoidea and Mactroidea, each
contain one brackish water species that can extend
into freshwater coastal drainages and so are included
here. In North America, the Dreissenacea is represented
by two introduced freshwater species and an estuarine
species. The Corbiculoidea includes 36 native and five
introduced species in six genera; the Unionoidea are
composed of 278 native species and 13 subspecies in 49
genera (Turgeon et al., 1998). Separate taxonomic keys
are provided here for the latter two supperfamilies.
1a.
Shell hinge ligament is external ..........................................................................................................................................................2
1b.
Shell hinge ligament is internal ...........................................................................................................................................................4
2a(1a).
Shell with lateral teeth extending anterior and posterior of true cardinal teeth (Fig. 1), shells of adults generally small (25 mm
in shell length, shell thin and fragile; exceptions are the genera Polymesoda and Corbicula)..............................................superfamily
Corbiculoidea [See Section V.B.]
2b.
Shell without lateral teeth extending anterior and posterior of cardinal teeth .....................................................................................3
3a(2b).
Shell hinge with two cardinal teeth and without lateral teeth; shell thin and fragile, 12 – 15 mm long with small umbos; Cyrenoida
floridana (Dall) (extends from brackish into coastal freshwater drainages in Florida) .....................................superfamily Cyrenoidea
3b.
Shell without true cardinal teeth, when present, lateral teeth only occur posterior to usually well-developed pseudocardinal
teeth (Fig. 1), pseudocardial teeth absent or vestigial in some species; Shells of adults are generally large (25 mm in shell
length) ................................................................................................................................superfamily Unionoidea [See Section V.C.]
4a(1b).
Hinge with anterior and posterior latoral teeth on either side of cardinals; Shell massive, adults 25 – 60 mm shell length, obliquely
ovate; Rangia cuneata (Gray) (extends from brackish into coastal freshwater drainages from Delaware to Florida to Veracruz,
Mexico) ..........................................................................................................................................................superfamily Mactroidea
4b.
Hinge without teeth; shell mytiloid in shape, anterior end reduced and pointed, hinge at anterior end, posterior portion of shell
expanded, anterior adductor muscle attached to internal apical shell septum, attached to hard substrates by byssal
threads .....................................................................................................................................................superfamily Dreissenoidea 5
5a(4b).
Periostracum bluish brown to tan without a series of dorsoventrally oriented black zigzag markings, anterior end hooked sharply
ventrally, ventral shell margins not distinctly flattened over entire ventral side of valves; restricted to brackish water habitats.
Mytilopsis leucophaeata (Conrad) (extends into coastal freshwater drainages from New York to Florida to Texas and Mexico)
............................................................................................................................................................................................Mytilopsis
5b.
Periostracum light tan and often marked with a distinct series of black vertical zigzag markings; anterior portion of shell not ventrally hooked, ventral shell margins can be flattened, restricted to freshwaters, Dreissena polymorpha (Pallas) (Fig. 24); (a European
species introduced into the Great Lakes in Lake St. Clair and present by December 2000 throughout the Great Lakes, the St.
Lawrence River and the Mississippi Drainage. and Dreissona bugensis Andrusov now in lakes Erie and Ontario, the Erie-Barge
Canal and the St. Lawrence River .........................................................................................................................................Dreissena
FIGURE 24
The external morphology of the shell of the
zebra mussel, Dreissena polymorpha.
398
Robert F. McMahon and Arthur E. Bogan
B. Taxonomic Key to the Genera of Freshwater
Corbiculacea
This key is based on the excellent species key for
North American freshwater Corbiculoidea by Burch
(1975a) with additional material from Clarke (1973)
for the Canadian Interior Basin, and Mackie et. al.
(1980) for the Great Lakes. The Corbiculoidea have
ovate, subovate, or trigonal shells with lateral hinge
teeth anterior and posterior to the cardinal teeth. All
North American species expect Corbicula fluminea are
in the family Sphaeriidae. The family designation
Pisidiidae has also been commonly applied to this
group, but the International Commission of Zoological
Nomenclature (ICZN) placed the Spheariidae (Name
number 573) on the Offical List of Family Names
(Opinion 1331) in 1985; hence, Sphaeriidae is used as
the family designation in this key and the rest of the
chapter. In North America, the Sphaeriidae comprise
the dominant bivalve fauna in small, often ephemeral
ponds, lakes, and streams, the profundal portions of
lakes and in silty substrates. Identification is generally
based on shell morphology, but requires, in some cases,
soft tissue morphology.
1a.
Shells large (maximum adult shell length 25 mm), thick, lateral teeth serrated .................................................family Corbiculida 2
1b.
Shells generally small (maximum shell length 25 mm), thin, lateral teeth smooth ..............................................family Sphaeriidae 4
2a(1a).
Maximum adult shell length generally 50 mm, shell ornamented by distinct, concentric sulcations, anterior and posterior lateral
teeth with many fine serrations, simultaneous hermaphrodites, massive numbers of small (length 0.3 mm) developmental stages
(1000) incubated directly in inner demibranchs, released juveniles (5 mm SL) anchor to substratum with a single mucilaginous
byssal thread (Fig. 25)........................................................................................................................................................Corbicula 3
FIGURE 25
External morphology of the shell valves of the light-colored shell morph (Corbicula fluminea)
and dark-colored shell morph (Corbicula sp.) of the North American Corbicula species complex. (A) Right and
left valves of Corbicula sp. (dark-colored morph). (B) Right and left valves of C. fluminea (light-colored
morph). (C) Anterior view of the shell valves of C. fluminea (light-colored morph). (D) Anterior view of the
shell valves of Corbicula sp. (dark-colored morph). Note the distinguishing shell characteristics of these two
species. C. fluminea, the light-colored morph, which is widely distributed in North America (Fig. 11), has a
more nearly trigonal shell, taller umbos, a greater relative shell width and more widely spaced concentric sulcations than does Corbicula sp., the dark-colored morph, that is limited to spring-fed, alkaline, lotic habitats in
the southwestern United States (Fig. 11). The dark-colored shell morph also has a dark olive green-to-black periostracum and deep royal blue nacre, while the light-colored shell morph has a yellow green-to-light brown
periostracum and white-to-light blue or light purple nacre.
11. Mollusca: Bivalvia
399
2b.
Maximum adult shell length generally 50 mm, shell ornamentation of many fine, closely spaced concentric striations, embryos not
incubated in demibranchs, dioecious, periostracum deep brown in color, three cardinal teeth, estuarine, restricted to brackish waters
in the tidal portions of rivers. Polymesoda caroliniana (Bosc) (Virginia to northern Florida to Texas)...............................Polymesoda
3a(2a).
Shell nacre white with light blue, rose, or purple highlights, particularly at shell margin, muscle scars of same color intensity as rest
of nacre, periostracum yellow to yellow-green or brown with outer margins always yellow or yellow-green in healthy, growing specimens, shell trigonal to ovate, umbos inflated and distinctly raised above dorsal shell margin, shell length : shell height ratio ⬇1.06,
shell length : shell width ratio ⬇1.47, shell height : shell width ratio ⬇ 1.38, concentric shell sulcations widely spaced, 1.5 sulcations/mm shell height (Hillis and Patton, 1982); introduced in the early 1900s, it has spread throughout drainage systems of the
United States and coastal northern Mexico (Fig. 11), the “light-colored shell morph” of Corbicula or Asian clam (Fig. 25B, C).Corbicula fluminea (Müller)
3b.
Shell nacre uniformly royal blue to deep purple over entire internal surface, muscle scars more darkly pigmented than rest of nacre,
periostracum dark olive green to black, edges of valves in healthy, growing specimens not yellow or yellow-green, shell more ovate
and laterally compressed with umbos less inflated and less distinctly raised above the dorsal shell margin than in C. fluminea, shell
length : height ratio ⬇1.15, shell length : width ratio ⬇ 1.65, shell height : width ratio ⬇ 1.43, concentric shell sulcations narrowly
spaced, particularly at umbos, 2.1 sulcations/mm shell height (Hillis and Patton, 1982); introduced, distribution limited to highly
oligotrophic, permanent, spring-fed, calcium carbonate-rich streams in the southwestern United States (Britton and Morton, 1986);
(Fig. 11). Called the dark-colored shell morph of Corbicula, its taxonomic status is uncertain, electrophoretic (Hills and Patton,
1982; McLeod, 1986) and physiological evidence (Cleland et al., 1986) suggest it to be distinct from C. fluminea (Fig. 25A, D)
........................................................................................................................................................................................Corbicula sp.
4a(1b).
Both inhalant and exhalant mantle cavity siphons present and well developed, umbos lie anterior of center......................................5
4b.
Only exhalant mantle cavity siphon present, inhalant siphon either absent or formed as a slit in the posterior-ventral mantle edges,
umbos posterior of center, generally small, shell length 0.5 – 12 mm, embryos in inner demibranch held in thick-walled sacs, each
with individual chambers for embryos, no byssal gland, 24 species widely distributed in North Amercia; for species identifications
and distributions see Burch (1975a) (Fig. 26A) .......................................................................................subfamily Pisidiinae Pisidium
FIGURE 26
Diagrams of the external morphology of the left shell valve of species representative of the
North American genera of the freshwater bivalve family, Sphaeriidae. Size scaling bar is 1-mm long.
400
Robert F. McMahon and Arthur E. Bogan
5a(4a).
Inhalant and exhalant mantle cavity siphons partially fused, embryos incubated in inner demibranchs in thin-walled longitudinal
pouches, no byssal gland, shell with two cardinal teeth in each valve, without external mottling....................subfamily Sphaeriinae 6
5b.
Inhalant and exhalant siphons not fused, embryos develop in individual chambers formed between inner and outer lamellae of inner
demibranchs, functional byssal gland present, only one cardinal tooth in each shell valve, with external, mottled pigmentation,
Eupera cubensis (Prime) (Atlantic coastal plain drainages from southern Texas to North Carolina, Caribbean Islands) (Fig. 26B) ......
.................................................................................................................................................................subfamily Euperinae Eupera
6a(5a).
Shell sculptured with relatively coarse or widely spaced striae ( 8 striae/mm in middle of shell), shell relatively massive and strong,
Sphaerium simile (Say) (Southern Canada from New Brunswick to British Columbia, south from Virginia to Wyoming, S. striatinum (Lamarck) (Canada from New Brunswick to the upper Yukon River, throughout the United States, Mexico, and Central
America), S. fabale (Prime) (Southern Ontario to Georgia and Alabama) (Fig. 26C) ...........................................................Sphaerium
6b.
Shell relatively thin, often fragile, with many fine, narrowly spaced striae (12 striae/mm in middle of shell) ...................................7
7a(6b).
Shell of adults 8 mm in length .........................................................................................................................................................8
7b.
Shell of adults 8 mm in length .........................................................................................................................................................9
8a(7a).
Posterior valve margin at near right angle to dorsal margin, shells roughly rhomboidal, umbos large and distinctly elevated above
dorsal shell margin, Musculium partumeium (Say) (United States and southern Canada), M. transversum (Say) (Canada and the
United States east of the continental divide, extending into Mexico), M. securis (Prime) (Nova Scotia to British Columbia southwestern Northwest Territories in Canada, United States except for southwest) (Fig. 26D)..................................................Musculium
8b.
Posterior and dorsal margins rounded or forming an obtuse angle, shells ovate, Sphaerium corneum (Linnaeus) (introduced from
Europe, localities in southern Ontario and Lakes Champlain and Erie), S. nitidum Westerlund (distribution is holarctic, northern
Canada to northern United States), S. occidentale (Prime) (Canada from New Brunswick to southeastern Manitoba, northern
United States south to Florida, west to Utah and Colorado) (Fig. 26 E, F)...........................................................................Sphaerium
9a(7b).
Umbos large, distinctly elevated above the dorsal shell margin........................................................................................................10.
9b.
Umbos small, indistinctly elevated above the dorsal shell margin, Sphaerium corneum (Linnaeus), (introduced, localities in Ontario,
Lakes Champlain and Erie) (Fig. 26E) .................................................................................................................................Sphaerium
10a(9a).
Shell rounded, umbos not prominent, Sphaerium occidentale (Prime) (New Brunswick to southeastern Manitoba, northern United
States south to Florida in the east and Utah and Colorado in the west) (Fig. 26F) ...............................................................Sphaerium
10b.
Posterior end of shell truncate, shell rhomboidal, umbos prominent, Musculium lacustre (Müller) (From treeline in Canada south
throughout all but southwestern United States into central America) (Fig. 26G).................................................................Musculium
C. Taxonomic Key to the Genera of Freshwater
Unionoidea
The Unionoidea make up the large bivalve fauna
(shell length 25mm) of permanent freshwater lakes,
rivers, and ponds. North America has the richest and
most diverse unionoidean fauna in the world, including
278 species and 13 recognized subspecies in 49 genera
in the Unionidae and five species in two genera in the
Margaritiferidae. The taxonomy used here follows Turgeon et al. (1998) with the addition of Johnson (1998),
and Williams and Fradkin (1999). Unionoidean taxonomy remains very uncertain because intraspecific and
interpopulation variation often makes identification and
systematics difficult. Unionoidean bivalve shells lack
true cardinal teeth and, when present, lateral teeth occur only posterior to pseudocardinal teeth (Fig. 1). Figure 27 displays shell-shape outlines and external shell
ornamentations referred to in these taxonomic keys.
The key in the first edition of this chapter, as well as
many of the major keys to freshwater bivalves (Walker,
1918; Clench, 1959; Burch, 1975b; Clarke, 1973), relied
on the integrated use of important anatomical structures
and shell characters to identify unionoidean bivalves.
However, most field biologists do not have the luxury of
having a fully gravid female in hand for identification,
but more likely, a dead shell or valve. Several colleagues
have commented that one can only use a key containing
anatomical characters to key out a shell when you already know what the animal is. Several state keys (e.g.,
Parmalee, 1967; Watters, 1995; Oesch, 1984; Strayer
and Jirka, 1997) have provided keys to shells of species
occurring within the political boundaries of a particular
state. North America is considered to consist of those
river systems and lakes north of, and including, the Rio
Grande Basin and the rest of the boundary with Mexico.
This key is artificial and the key is divided into four sections corresponding to geographical provinces to facilitate identification. The four sections are: Gulf Coast, including Florida; Mississippi River Basin, including the
Interior Basin of Canada; the area west of the Rocky
Mountain divide; and the Atlantic Coast. These subdivisions are a simplification of the 12 faunal provinces recognized by Parmalee and Bogan (1998). Table VII lists
11. Mollusca: Bivalvia
401
FIGURE 27 Illustrations of the diagnostic shell features or characters used for
taxonomic identification in Section V.C. Shell-shape descriptions: (A) rhomboidal; (B)
triangular or trigonal; (C) round; (D) quadrate; (E and F) oval or ovoid; and (G) elliptical. Posterior shell-ridge morphology: (H) posterior ridge convex; and (I) posterior ridge
concave. Concentric ridge structures of umbos: (J) single-looped concentric ridges; (K)
double-looped concentric ridges; (L); coarse concentric ridges; and (M) fine concentric
ridges. (Redrawn from Burch, 1975b.)
the genera found in each of these four sections. This geographical approach allows a biologist to focus only on
those genera actually occurring in their region and ignores the rest of the diversity (e.g., west of the Rocky
Mountains has only three genera: one Margaritiferidae
and two Unionidae). We have attempted to provide a
logical and clear key to the unionoidean genera of North
America based solely on shell characters. The valve or
pair of valves to be identified is assumed for the purposes of this key to be adults. Identification of very small
juvenile shells and small adult specimens can be difficult.
Landmarks, structures and shell shape are based on
complete shells. We have attempted to use obvious characters, but many of them require familiarity with
unionoidean shell morphology and the range of varia-
tion in each of these characters. Strayer and Jirka (1997;
p. 28) stated the problem most clearly when they said,
“Keys based on shell characters are inevitably filled with
vague, subjective terms, are frustrating for beginners to
use, and misidentify many shells.” They have finally put
into print what most malacologists have said: “Users
should know that if they rely solely on this key, they will
misidentify many shells.” (Strayer and Jirka, 1997; p.
28). There is no substitute for comparing the specimen
to be identified with other identified specimens in a museum collection or specimens that have had their identification verified by a specialist. This key is an introduction
to the genera of North American unionoideans. Each
couplet which contains a genus, also lists some of the
species in that genus which should key out to that cou-
402
Robert F. McMahon and Arthur E. Bogan
TABLE VII
List of Unionoidean Genera by Geographic Area Used in the Key
Margaritiferidae
Pacific coast
Atlantic coast
Gulf coast
Margaritifera
Margaritifera
Margaritifera
Unionidae
Alasmidonta
Anodonta
Anodonta
Anodontoides
Alasmidonta
Amblema
Anodonta
Anodontoides
Arcidens
Interior basin
Cumberlandia
Actinonaias
Alasmidonta
Amblema
Anodonta
Anodontoides
Arcidens
Arkansia
Cyclonaias
Cyprogenia
Cyrtonaias
Disconaias
Ellipsaria
Elliptio
Elliptoideus
Epioblasma
Fusconaia
Glebula
Elliptio
Fusconaia
Dromus
Ellipsaria
Elliptio
Epioblasma
Fusconaia
Gonidea
Lampsilis
Lasmigona
Lampsilis
Lasmigona
Leptodea
Lexingtonia
Ligumia
Leptodea
Ligumia
Medionidus
Megalonaias
Obliquaria
Obovaria
Plectomerus
Pleurobema
Uniomerus
Utterbackia
Pleurobema
Popenaias
Potamilus
Ptychobranchus
Pyganodon
Quadrula
Quincuncina
Strophitus
Toxolasma
Tritogonia
Truncilla
Uniomerus
Utterbackia
Villosa
18
Villosa
37
Pyganodon
Strophitus
Toxolasma
Total
3
plet. This list of species is not complete, but is representative of the species in the genus with the listed set of
characters. Two genera are a source of much confusion,
Elliptio and Pleurobema. The South Atlantic Slope Elliptio species complex has yet to be worked out. Johnson (1970) lumped a considerable number of taxa,
Hemistena
Lampsilis
Lasmigona
Lemiox
Leptodea
Lexingtonia
Ligumia
Medionidus
Megalonaias
Obliquaria
Obovaria
Pegias
Plectomerus
Plethobasus
Pleurobema
Potamilus
Ptychobranchus
Pyganodon
Quadrula
Simpsonaias
Strophitus
Toxolasma
Tritogonia
Truncilla
Uniomerus
Utterbackia
Venustaconcha
Villosa
43
which may, in fact, be valid species. The Gulf Coast
Pleurobema species complex is the other major area of
confusion. There are a large number of named taxa, but
it is not clear at the present time which is a valid species
and which is part of a cline. This group is still in need of
further work. We would recommend that, to identify
11. Mollusca: Bivalvia
specimens to the species level, the user move to a key for
the state or province in which he/she is working. These
volumes will give detailed descriptions, keys and figures
of the species occurring in your local area (e.g., Florida:
Clench and Turner, 1956; Johnson, 1972; Tennessee:
403
Parmalee and Bogan, 1998; Louisiana: Vidrine, 1993;
New York: Strayer and Jirka, 1997; Illinois: Parmalee,
1967; Missouri: Oesch, 1984; and Ohio: Watters, 1995).
A detailed list by state of freshwater bivalve literature
can be found in Williams et al. (1993).
1a.
Origin of shell is from west of the Rocky Mountains ...................................................................................................................4
1b.
Shell is not from this area .............................................................................................................................................................2
2a. (1b)
Origin of shell is from the Atlantic Coast of North America ........................................................................................................6
2b.
Shell is not from this area .............................................................................................................................................................3
3a. (2b)
Origin of shell is from the Mississippi River Basin......................................................................................................................26
3b.
Origin of shell is from the Gulf Coast of North America............................................................................................................89
West of the Rocky Mountains
4a. (1a)
Shell has lateral muscle scars [small pits for the attachment of mantle muscles to the shell], typically dark periostracum, relatively thick shell with pseudocardinal and lateral teeth .............................................................Margaritifera falcata (Fig. 11.28B)
4b.
Shell lacks lateral muscle scars, periostracum typically not dark and hinge teeth reduced or absent .............................................5
5a. (4b)
Shell has a very strongly developed posterior ridge, reduced hinge teeth, occurs from British Columbia to central California, east
to Nevada and Idaho .........................................................................................................................Gonidea angulata (Fig. 28U)
5b.
Shell lacks the sharp posterior ridge, lacks any evidence of hinge teeth .....................................Anodonta [in part] [A. beringiana,
A. californiensis, A. dejecta, A. kennerlyi, A. nuttalliana, A. oregonensis] (Fig. 28F)
Atlantic Coast
6a. (2a)
Shell has lateral muscle scars, typically dark periostracum, relatively thick shell with pseudocardinal teeth and lateral teeth .........
............................................................................................................................................Margaritifera margaritifera (Fig. 28B)
6b.
Shell lacks lateral muscle scars, periostracum may not be dark, hinge teeth present or absent ......................................................7
7a. (6b)
Shell with hinge teeth absent or greatly reduced ...........................................................................................................................8
7b.
Shell with pseudocardinal teeth present, with or without lateral teeth ........................................................................................12
8a. (7a)
Umbo not projecting above the hinge-line ...........................................................Utterbackia [in part] [U. imbecillis] (Fig. 28AW)
8b.
Umbo projecting above the hinge-line ..........................................................................................................................................9
9a. (8b)
Beak sculpture double looped (Fig. 27), shell uniformly thin..................................Pyganodon [in part] [P. cataracta] (Fig. 28AN)
9b.
Beak sculpture consists of concentric bars ..................................................................................................................................10
10a. (9b)
Nacre usually orange in the beak cavity, pseudocardinal tooth area represented by a thickening near the umbo, ventral shell
margin uniform thickness .......................................................................................Strophitus [in part] [S. undulatus] (Fig. 28AR)
10b.
Nacre bluish or white, hinge plate uniformly thin, teeth or swellings absent, ............................................................................11
11a. (10b)
Ventral margin with a prominent thickened area along the anterior ventral magin below the pallial line, Anodonta [in part] [A.
implicata] (Fig. 28F)
11b.
Ventral margin not as above, shell elongate, Anodontoides [in part] [A. ferussacianus, restricted to the Susquehanna and Hudson River basins] (Fig. 28G)
12a. (7b).
Shell with lateral teeth absent or reduced, neither functional nor interlocking ..............................................Alasmidonta [in part]
[A. varicosa, A. marginata susquehannae, A. undulata] (Fig. 28D)
12b.
Shell truncated, with well-developed lateral teeth .......................................................................................................................13
13a. (12b)
Right valve with two lateral teeth, small, rare ......................................................Alasmidonta [in part] [A. heterodon] (Fig. 28D)
13b.
Right valve with one lateral tooth ..............................................................................................................................................14
14a. (13b)
Shell with spines on the umbo and down on to the disk of the shell ...........................................................................................15
14b.
Shell lacks any evidence of spines ...............................................................................................................................................16
15a. (14a)
Shell thick and may be large .............................Elliptio [in part] [shell from the Altamaha River Basin, Georgia, Elliptio spinosa,
or the Tar/Neuse River Basin, North Carolina Elliptio steinstansana] (Fig. 28P)
15b.
Shell small and from the James River Basin, Virginia .....................................................................Pleurobema collina (Fig. 28AJ)
404
Robert F. McMahon and Arthur E. Bogan
405
FIGURE 28 Diagrams of the external morphology mostly of right valves of the shell of species representative
of the North American genera of the freshwater bivalve superfamily, Unionoidea (Figs. 28A – AY). Figures
B – E, G – K, N, P, R – AC, AE – AG, AI, AJ, AL – AQ, AR – AU, reprinted with permission of Mrs. W.T.
Edmondson from W.T. Edmondson (1959); Figures Q, AW, reprinted with permission of J.B. Burch from Burch
(1973); Figures A, F, L, M, AD, AH, AK, AX, AY, reprinted with permission from the Treatise on Invertebrate
Paleontology, courtesy of The Geological Society of America and the University of Kansas, © 1969.
(Continues)
406
Robert F. McMahon and Arthur E. Bogan
FIGURE 28
(Continued)
11. Mollusca: Bivalvia
FIGURE 28
(Continued)
407
408
Robert F. McMahon and Arthur E. Bogan
16a. (14b)
Hinge line in left valve with an additional small interdental or accessory tooth, giving the appearance of three pseudocardinal
teeth, shell more or less compressed, shell shape rhomboid, periostracum dark green with numerous green rays, beak sculpture
consists of prominent bars ...................................................Lasmigona [in part][L. decorata, L. robusta, L. subviridis](Fig. 28X)
16b.
Left valve without extra interdental tooth ..................................................................................................................................17
17a. (16b)
Shell shape rectangular to broadly triangular .............................................................................................................................18
17b.
Shell shape oval, round or rhomboid..........................................................................................................................................19
18a. (17a)
Shell is from the James River Basin, Virginia .............................................................................Lexingtonia subplana (Fig. 28AA)
18b.
Shell is from an area extending from the Roanoke River Basin south to the headwaters of the Savannah River Basin....................
...........................................................................................................................................................Fusconaia masoni (Fig. 28S)
19a. (17b)
Shell shape rhomboid or rectangular ..........................................................................................................................................20
19b.
Shell shape oval or round ...........................................................................................................................................................23
20a. (19a)
Shell usually more than twice as long as high .............................................................................................................................21
20b.
Shell usually less than twice as long as high................................................................................................................................25
21a. (20a)
Nacre color white, shell inflated, ...............................................................................................................................................22
21b.
Nacre color typically some shade of purple, but ranges from white to salmon to purple .......................................Elliptio [in part]
[this genus contains basically three shell shapes, narrow and elongate: the Elliptio lanceolata complex; rectangular with various
degrees of inflation: the Elliptio complanata complex; those shells with short shell length, not too tall and inflated: the Elliptio
icterina complex] (Fig. 28P)
22a. (21a)
Periostracum unrayed, shell thick, posterior end angled, periostracum mat or fuzzy, rectangular in shell shape, Uniomerus caroliniana (Fig. 28 AV)
22b.
Periostracum rayed in juveniles, posterior end tapered to a point in middle of posterior margin, periostracum not mat, Ligumia
nasuta (Fig. 28AB)
23a. (19b)
Adult shell typically 40 mm in length, with a fuzzy or mat textured dark periostracum ...............................Toxolasma [in part]
[T. pullus] (Fig. 28AS)
23b.
Adult shell 40 mm in length, lacking the pronounced fuzzy periostracum ...............................................................................24
24a. (23b)
Shell shape oval to elongate oval, periostracum very shiny to mat with rays ...................................Lampsilis [in part] [L. cariosa,
L. dolabraeformis, L. radiata, L. splendida] (Fig. 28W)
24b.
Shell shape oval, periostracum dull yellow, without rays or with fine rays all over the shell, found in or near tidewater, nacre often a salmon color .......................................................................................................Leptodea [in part] [L. ochracea] (Fig. 28Z)
25a. (20b)
Periostracum light colored with numerous green rays, shell relatively thin, oval to elongate oval, bladelike pseudocardinal teeth
.......................................................................................................Villosa [in part] [V. delumbis, V. villosa, V. vibex] (Fig. 28AY)
25b.
Periostracum dark to black, shell thick, no green rays, shell shape oval to round...................................................Villosa [in part]
[V. constricta] (Fig. 28AY)
Mississippi River Basin
26a. (3a)
Shell with lateral muscle scars on the inside center of the valve ............................................Cumberlandia monodonta (Fig. 28A)
26b.
Shell lacking any evidence of lateral muscle scars .......................................................................................................................27
27a. (26b)
Shell not as above, lacking lateral teeth or all evidence of any hinge teeth ..................................................................................28
27b.
Shell with both pseudocardinal and lateral teeth present ............................................................................................................34
28a. (27a)
Shell with pseudocardinal teeth, but lacking lateral teeth ...........................................................................................................29
28b.
Shell with greatly reduced or totally lacking both pseudocardinal and lateral teeth ....................................................................30
29a. (28a)
Shell 25 mm, typically periostracum eroded off with periostracum restricted to narrow strip on shell margin, beak sculpture
heavy bars extending down on the disk of the shell, restricted to the upper Cumberland and upper Tennessee River basins ..........
...............................................................................................................................................................Pegias fabula (Fig. 28AG)
29b.
Shell not small, periostracum not eroded or erosion restricted to the umbo, beak sculpture consists of heavy bars, widespread in
the Mississippi River Basin..................................................................Lasmigona [in part] [L. costata, L. complanata] (Fig. 28X)
30a. (28b)
Shell with pseudocardinal tooth consisting of a slight swelling or knob, lateral tooth consisting of a rounded ridge, beak cavity
copper colored, umbo centrally located ......................................................................................Strophitus undulatus (Fig. 28AR)
30b.
Hinge teeth completely absent ....................................................................................................................................................31
31a. (30b)
Umbonal area of shell projects above the plane of the hinge line ................................................................................................32
31b.
Umbo level with hinge line or below the level of the hinge line ..................................................................................................33
11. Mollusca: Bivalvia
409
32a. (31a)
Umbo elevated well above the hinge line, shell shape variable, ranging from oval, elliptical to rhomboid, beak sculpture consists
of double loopedridges (Fig. 27) with projections on the bottom of the loop ...........Pyganodon [in part] [P. grandis] (Fig. 28AN)
32b.
Umbo projecting only slightly above the hinge line, shell elongated, inflated, beak sculpture consists of 3 or 4 fine sharp concentric ridges ...........................................................................................................................Anodontoides ferussacianus (Fig. 28G)
33a. (31b)
Shell shape round to circular, compressed, beak sculpture consists of 4 – 5 pairs of broken irregular double looped ridges.............
............................................................................................................................Anodonta [in part] [A. suborbiculata] (Fig. 28F)
33b.
Shell shape elongate, inflated, dorsal and ventral margins nearly straight and parallel, beak sculpture consists of 5 – 6 fine irregular concentric ridges ............................................................................................Utterbackia [in part] [U. imbecillis] (Fig. 28AW)
34a. (27b)
Shell with sculpture on the external shell surface........................................................................................................................35
34b.
Shell with no plications, undulations, ridges, pustules or nodules...............................................................................................55
35a. (34a)
Shell with prominent pustules or nodules and/or undulations, plications or ridges .....................................................................36
35b.
Shell with only prominent, well-defined pustules, knobs or nodules ...........................................................................................46
36a. (35a)
Shell with undulations, ridges, or plications, no pustules or nodules on the surface....................................................................37
36b.
Shell with undulations, ridges or plications and pustules on the disk or umbonal area ...............................................................44
37a. (36a)
Shell with undulations, plications, ridges restricted to the posterior slope .................................................................................38
37b.
Shell with undulations, plications, ridges not restricted to the posterior slope ............................................................................41
38a. (37a)
Shell shape rectangular to oval, slablike, compressed, left valve with an accessory dentacle posterior to pseudocardinal teeth .......
.....................................................................................................................................Lasmigona [in part] [L. costata](Fig. 28X)
38b.
Shell shape more elongate, lacks the accessory dentacle..............................................................................................................39
39a. (38b)
Shell elongate, inflated with a blue to blue-green nacre color, restricted to the Tennessee and Cumberland River basins ................
..........................................................................................................................Medionidus [in part] [M. conradicus] (Fig. 28AC)
39b.
Shell elongate, inflated, nacre not as above.................................................................................................................................40
40a. (39b)
Shell thin, posterior slope steeply angled to moderately angled ............................Alasmidonta [in part] [A. marginata] (Fig. 28D)
40b.
Shell thicker, posterior slope rounded, posterior ridge rounded, restricted to the Tennessee and Cumberland River Basins ............
....................................................................................................................Ptychobranchus [in part] [P. subtentum] (Fig. 28AM)
41a. (37b)
Shell shape rectangular, with a distinct posterior ridge, purple nacre, open and shallow beak cavity ..............................................
...........................................................................................................................................Plectomerus dombeyanus (Fig. 28AH)
41b.
Shell without a distinct posterior ridge, typically white nacre .....................................................................................................42
42a. (41b)
Shell thick, compressed, maximum shell length 50 mm, low plications or wrinkles restricted to the posterior slope and posterior ventral margin of the shell, this species restricted to the Tennessee River Basin..............................Lemiox rimosus (Fig. 28Y)
42b.
Shell not as above.......................................................................................................................................................................43
43a. (42b)
Shell variably inflated, shell shape round to quadrate, heavy pseudocardinal teeth, deep compressed beak cavity, sculpture consists of typically three well-developed ridges or undulations running across the shell from anterior of the umbo to the posterior
ridge, widespread in the Mississippi River Basin..................................................................................Amblema plicata (Fig. 28E)
43b.
Shell somewhat inflated, shell shape quadrate, or approaching oval, pseudocardinal teeth curved, heavy sculpturing on the posterior half of the shell, species is restricted to the Red River Drainage in Oklahoma and the Ouachita River Drainage in
Arkansas .............................................................................................................................................Arkansia wheeleri (Fig. 28I)
44a. (36b)
Shell elongate with a well-developed diagonal posterior ridge .....................................................Tritogonia verrucosa (Fig. 28AT)
44b.
Shell shape not elongate, lacking well-defined posterior ridge.....................................................................................................45
45a. (44b)
Well-developed plications running across the shell and pustules on the disk and umbonal area ..................................Megalonaias
nervosa (Fig. 28AD)
45b.
Plications not well developed and wavy, not straight, plications on posterior slope, two rows of pustules or knobs on umbo........
......................................................................................................................................................Arcidens confragosa (Fig. 28H)
46a. (35b)
Pustules distributed across most of the shell ...............................................................................................................................47
46b.
Pustules restricted to a single or double row...............................................................................................................................50
47a. (46a)
Pustules covering most of the shell, purple nacre, beak cavity deep and compressed .................. Cyclonaias tuberculata (Fig. 28J)
47b.
Pustules covering most of the shell, nacre color white to pink not purple ...................................................................................48
48a. (47b)
Pustules consistently small, with a shallow sulcus down the center of the disk, nacre white ...........................Cyprogenia [in part]
[C. stegaria in the Ohio, Tennessee and Cumberland rivers; C. aberti in Arkansas, Oklahoma] (Fig. 28K)
48b.
Pustules generally larger, and often of variable size and shape ....................................................................................................49
410
Robert F. McMahon and Arthur E. Bogan
49a. (48b)
Disk of the shell usually with a broad green stripe running down the disk of the shell with a variable number of pustules, varies
from a very few to covering most of the shell, nacre color mostly white, beak cavity deep and open not compressed ....................
...............................................................................................................................Quadrula [in part] [Q. pustulosa] (Fig. 28AO)
49b.
Disk of shell without a broad green stripe, consistently covered with a large number of pustules, nacre color often pink, salmon
or white, beak cavity open and shallow..............................................................Plethobasus [in part] [P. cooperianus] (Fig. 28AI)
50a. (46b)
Pustules forming two distinct rows.............................................................................................................................................51
50b.
Pustules or nodules forming a single row....................................................................................................................................52
51a. (50a)
Two rows of pustules with one row on the posterior ridge, shell thick ......Quadrula [in part], [Q apiculata, this species has small
pustules in the sulcus between the two rows of pustules; Q. nodulata, this species lacks a sulcus between the rows of pustules
and may only have very few pustules; Q. quadrula; Q. fragosa, this species has a well-developed wing posterior to the umbo]
(Fig. 28AO)
51b.
Radiating row of knobs, shell inflated, thin to relatively thick, umbo high and full, pseudocardinal teeth compressed, not
curved; beak sculpture consists of irregular nodules forming two loops which continue down and across most of the shell in two
radiating rows, [mostly in small and juvenile shells] pustules fuse, become elongate and form plications, plications on the posterior slope of adults .......................................................................................................................Arcidens confragosus (Fig. 28H)
52a. (50b)
The single row of pustules or knobs restricted to the posterior ridge .................................................................Quadrula [in part]
[Q. cylindrica, with pustules on the posterior ridge while Q. cylindrica strigillata of the upper Tennessee River Basin has small
pustules over most of the shell as well as pustules on the posterior ridge.] (Fig. 28AO)
52b.
The single row of pustules or knobs forming a single row down the center of the disk...............................................................53
53a. (52b)
Pustules large, located in a row down the center of the shell and limited to about three pustules on each valve .............................
.......................................................................................................................................................Obliquaria reflexa (Fig. 28AE)
53b.
Pustules more numerous, but still in a centrally located row ......................................................................................................54
54a. (53b)
Pustules usually wider than tall, shell periostracum a waxy yellow color .......................................................Plethobasus [in part]
[P. cicatricosus, P. cyphyus] (Fig. 28AI)
54b.
Pustules rather small, running down the centerline of the disk, but interrupted by a well-defined, raised ridge running around
the shell like a growth line, restricted to the Tennessee and Cumberland River drainages ....................Dromus dromas (Fig. 28N)
55a. (34b)
Shell with a well-developed dorsal wing projecting above the hinge line, the wing usually posterior of the umbo, but some
species may also have an anterior wing [some older shells, such as in Leptodea fragilis, maybe lacking the dorsal wing] ...........56
55b.
Shell without a well-developed dorsal wing ................................................................................................................................58
56a. (55a)
Hinge teeth thin pseudocardinal teeth compressed, thin and not projecting perpendicular to the axis of the hinge line ..................
...................................................................................................................Leptodea [in part] [L. fragilis, L. leptodon] (Fig. 28Z)
56b.
Well-developed dorsal wing ........................................................................................................................................................57
57a. (56b)
White nacre, thick-shelled ....................................................................................Lasmigona [in part] [L. complanata] (Fig. 28X)
57b.
Purple nacre, thin-to-thick shelled, may be inflated.................................Potamilus [in part] [P. alatus, P. purpuratus] (Fig. 28AL)
58a. (55b)
Shell shape round .......................................................................................................................................................................59
58b.
Shell shape not round .................................................................................................................................................................68
59a. (58a)
Shell compressed, shell shape round ...........................................................................................................................................60
59b.
Shell not compressed, but more inflated, shell shape round, oval................................................................................................61
60a. (59a)
Shell compressed, shape nearly circular, without lateral teeth ...............................Lasmigona [in part] [L. complanata] (Fig. 28X)
60b.
Shell compressed, with lateral teeth, steep posterior slope, shell thick with well-developed hinge teeth, periostracum light yellow
with interrupted rows of green chevrons ..........................................................................................Ellipsaria lineolata (Fig. 28O)
61a. (59b)
Umbo central or nearly central in the dorsal margin...................................................................................................................62
61b.
Umbo anterior of the center of the shell .....................................................................................................................................65
62a. (61a)
Broad green ray extending from the umbo down only a short distance onto the disk of the shell ......................Quadrula [in part]
[Q. pustulosa without pustules] (Fig. 28AO)
62b.
No broad green ray ...................................................................................................................................................................63
63a. (62b)
Shell with a raised ridge one quarter of the distance from the umbo running around the shell parallel with the growth lines. Restricted to the Tennessee and Cumberland River basins .......................................................................Dromus dromas (Fig. 28N)
63b.
Shell without a raised ridge ........................................................................................................................................................64
11. Mollusca: Bivalvia
411
64a. (63b)
Shell surface smooth, without a broad central sulcus ..............................................................Obovaria [in part] [O. jacksoniana,
from the Mississippi River embayment shell somewhat more oval than round; O. retusa, nacre color white; nacre color purple
inside the pallial line, white outside of the pallial line; O. subrotunda, shell round, thick shelled, nacre typically white but some
populations with deep purple.] (Fig. 28AF)
64b.
Shell with broad shallow-to-pronounced sulcus, fine green rays, shallow beak cavity, well-developed hinge teeth, female shells
with a swollen, extended or expanded portion of the posterior slope and posterior ventral margin of the shell..............................
Epioblasma [in part] [E. capsaeformis, E. florentina florentina, E. florentina walkeri, E. lewisi, E. flexuosa, E. biemarginata, E.
stewardsoni, E. propinqua, E. torulosa torulosa, E. torulosa gubernaculum, E. torulosa biloba, E. sampsoni] (Fig. 28R)
65a. (61b)
Beak cavity deep compressed ......................................................................Fusconaia [in part][F. subrotunda, F. ebena] (Fig. 28S)
65b.
Beak cavity shallow and open.....................................................................................................................................................66
66a. (65b)
Restricted to the Tennessee River Basin, shell with green rays at least on the umbo area with a thick shell, heavy lateral teeth,
very shallow sulcus or missing, shallow-to-no beak cavity ..................................................Lexingtonia dolabelloides (Fig. 28AA)
66b.
Not restricted to the Tennessee River Basin, but widespread ......................................................................................................67
67a. (66b)
Long axis of the main pseudocardinal teeth parallel with the lateral teeth, umbo anterior ..............................................Obovaria
[in part, O. olivaria] (Fig. 28AF)
67b.
Teeth not as above, shell wedge shaped to triangular or approaching square with or without a central sulcus, nacre white to
deep pink ................................................................................................Pleurobema [in part] [P. rubrum, P. sintoxia] (Fig. 28AJ)
68a. (58b)
Shell shape not oval....................................................................................................................................................................69
68b.
Shell shape oval to oblong ..........................................................................................................................................................85
69a. (68a)
Shell shape rectangular, triangular, square with sulcus................................................................................................................70
69b.
Shell shape elongate, two-to-four times longer than high............................................................................................................75
70a. (69a)
Beak cavity deep.........................................................................................................................................................................71
70b.
Beak cavity shallow or broad and open ......................................................................................................................................72
71a. (70a)
Shell shape rectangular, with green rays on the umbo .......................................................................................Fusconaia [in part]
[F. flava, F. cuneolus, F. cor, F. ozarkensis, F. barnesiana has a shallow beak cavity but belongs with this group] (Fig. 28S)
71b.
Shell shape triangular to broadly triangular..........................................Pleurobema [in part] [P. cordatum, P. plenum] (Fig. 28AJ)
72a. (70b)
Posterior slope not very sharp or rounded, covered with lines of green chevrons ................................................Truncilla [in part;
T. truncata, T. donaciformis] (Fig. 28AU)
72b.
Posterior slope very sharp or steep .............................................................................................................................................73
73a. (72b)
Shell thin, inflated, abruptly truncate posteriorly, posterior slope lighter colored than the rest of the shell .....................................
Alasmidonta [in part] [A. marginata; shell thin, shell shape elongate, posterior slope not a steep sharp angle, headwaters of the
Little Tennessee River in western North Carolina, A. raveneliana; streams of the upper Cumberland River Basin, A. atropurpurea] (Fig. 28D)
73b.
Shell thin to thick, inflated posterior slope steep shell thick, shell shape rectangular...................................................................74
74a. (73b)
Females with posterior slope inflated into a marsupial swelling, males evenly rounded posterior ridge and not inflated posterior
ridge, nacre white ........................................................Epioblasma [in part] [E. triquetra wide spread, E. arcaeformis extinct and
formerly restricted to the Tennessee and Cumberland River basins.] (Fig. 28R)
74b.
Posterior margin not modified into a marsupial swelling, nacre purple..........................Elliptio [in part] [E. crassidens] (Fig. 28P)
75a. (69b).
Thin shelled................................................................................................................................................................................76
75b.
Thick shelled ..............................................................................................................................................................................79
76a. (75a)
Shell compressed, very thin, periostracum rayed, nacre iridescent except for a purple wash in the open beak cavity ......................
...............................................................................................................................................................Hemistena lata (Fig. 28V)
76b.
Shell inflated...............................................................................................................................................................................77
77a. (76b)
Hinge teeth thin, periostracum rayless, adult shell length 50 mm ...........................................Simpsonaias ambigua (Fig. 28AQ)
77b.
Periostracum usually rayed, adult shell length 50 mm .............................................................................................................78
78a. (77b)
Shell hinge thin, usually heavily rayed....................................................Villosa [in part] [V. iris complex, V. taeniata] (Fig. 28AY)
78b.
Shell yellow, plain or rayed ........................Lampsilis [in part] [L. teres, L. rafinesqueana, L. siliquoidea, L. virescens] (Fig. 28W)
79a. (75b)
Shell compressed ........................................................................................................................................................................80
79b.
Shell inflated...............................................................................................................................................................................82
412
Robert F. McMahon and Arthur E. Bogan
80a. (79a)
Adult shell length 40 mm, rayed, compressed, widespread, ...........................................Villosa [in part] [V. fabalis] (Fig. 28AY)
80b.
Adult shell length 40 mm ........................................................................................................................................................81
81a. (80b)
Shell hinge massive, curved, nacre white, periostracum with interrupted green rays ................................Ptychobranchus [in part]
[P. fasciolaris, P. occidentalis] (Fig. 28AM)
81b.
Hinge lighter, straight, nacre purple to white, periostracum typically unrayed ..................Elliptio [in part] [E. dilatata] (Fig. 28P)
82a. (79b).
Adult shell length 55mm
82b.
Adult shell length 55mm
83a. (82a).
Periostracum olive green with numerous rays Villosa [in part] [V. trabalis, restricted to the Tennessee River Basin above Muscle
Shoals and the upper Cumberland River basin, inflated, white nacre, V. perpurpurea, upper Tennessee River Basin, inflated, purple nacre] (Fig. 28AY)
83b.
Periostracum ranges from greenish to black without rays Toxolasma [in part] [T. lividus, purple nacre, T. parvus white nacre, T.
texasiensis white to salmon nacre] (Fig. 28AS)
84a. (82b)
Shell inflated, periostracum rayless, posterior shell margin comes to a blunt point ventrally, coarse concentric beak sculpture
opening posteriorly .......................................................................................Uniomerus [U. tetralasmus, U. declivus] (Fig. 28AV)
84b.
Shell inflated, periostracum rayed as a juvenile, becoming dark brown to black as an adult, posterior shell margin comes to a
point in middle of posterior margin, nacre white to white with purple wash in umbo area ...............................................Ligumia
[L. nasuta, L. recta, L. subrostrata] (Fig. 28AB)
85a. (68b)
Shell shape oval ..........................................................................................................................................................................86
85b.
Shell shape oblong......................................................................................................................................................................87
86a. (85a)
Shell shape oval, inflated, rayed, thin to thick shell, posterior ridge rounded or sharp, nacre variously white to pink or salmon
..............Lampsilis [in part] [L. cardium, L. fasciola, L. ovata, L. reeviana, L. rafinesqueana, L. higginsi, L. abrupta] (Fig. 28W)
86b.
Shell shape elongate oval ............................................Villosa [in part] [V. taeniata, interrupted broad green rays, restricted to the
Tennessee and Cumberland River basins; in the following three species males are oval while female shells have a truncate posterior margin, V. vanuxemensis, purple nacre, restricted to the Tennessee River Basin and the central Cumberland River Basin, V.
ortmanni, restricted to Kentucky, V. lienosa, variously colored nacre] (Fig. 28AY)
87a. (85b)
Shell shape oblong, inflated to globose, unrayed...................................Potamilus [in part] [S-shaped, very compressed hinge line,
very inflated and thin shelled P. capax; inflated, black shiny periostracum and purple nacre P. purpuratus] (Fig. 28AL)
87b.
Shell shape oblong, inflated but not globose ...............................................................................................................................88
88a. (87b)
Adult shell length 70 mm, shell shape elliptic to oblong, thick shelled, nacre white with a tinge of salmon to coppery color in
center of the shell, shallow beak cavity, rayed, with those rays on the posterior slope thick and wavy ....................Venustaconcha
ellipsiformis (Fig. 28AX)
88b.
Adult shell length 70 mm, shell shape oblong to oval, thin to thick shelled, nacre white to iridescent, beak cavity broad and
open, moderately deep, usually rayed but rays on posterior slope not thin or wavy ..............................Actinonaias [A. pectorosa,
thin shelled, restricted to the Tennessee and Cumberland River basins; A. ligamentina, widespread and typically thick shelled.]
(Fig. 28C)
Gulf Coast
89a (3b)
Shell with lateral muscle scars in the interior center of the valve ..................................................................Margaritifera [in part]
[M. hembeli, M. marrianae] (Fig. 28B)
89b.
Shell without lateral muscle scars in the interior center of the valve ...........................................................................................90
90a. (89b)
Shell with pseudocardinal and lateral teeth greatly reduced or absent ........................................................................................91
90b.
Shell with both pseudocardinal and lateral teeth present ............................................................................................................95
91a. (90a)
Pseudocardinal and lateral teeth present, but reduced ................................................................................................................92
91b.
Pseudocardinal and lateral teeth completely absent ....................................................................................................................93
92a. (91a)
Shell with pseudocardinal tooth consisting of a slight swelling or knob, lateral tooth consisting of a rounded ridge, beak cavity
copper colored, umbo centrally located ....................................Strophitus [in part] [S. subvexus, S. connasaugaensis] (Fig. 28AR)
92b.
Shell with pseudocardinal tooth thin, reduced and compressed, umbo projecting only slightly above the hinge line, shell elongated, inflated, beak sculpture consists of 3 – 4 fine, sharp, concentric ridges ............................................................Anodontoides
[in part] [A. radiatus] (Fig. 28G)
93a. (91b)
Umbo elevated well above the hinge line, shell shape variable, ranging from oval, elliptic or rhomboid, beak sculpture consists
of double looped with projections on the bottom of the loop ...................................Pyganodon [in part] [P. grandis] (Fig. 28AN)
93b.
Umbo level ridges with hinge line or below the level of the hinge line ........................................................................................94
94a. (93b)
Shell shape round to circular, compressed, beak sculpture consists of 4 – 5 pairs of broken irregular double looped ridges.............
............................................................................................................Anodonta [in part] [A. suborbiculata, A. heardi] (Fig. 28F)
11. Mollusca: Bivalvia
413
94b.
Shell shape elongate, inflated dorsal and ventral margins nearly straight and parallel, beak sculpture consists of 5 – 6 fine irregular concentric ridges................................................Utterbackia [in part] [U. imbecillis, U. peggyae, U. peninsularis] (Fig. 28AW)
95a. (90b)
Shell with sculpture on the external shell surface........................................................................................................................96
95b.
Shell without plications, undulations, ridges, pustules or nodules.............................................................................................109
96a. (95a)
Shell with pustules or nodules and/or undulations, plications or ridges ......................................................................................97
96b.
Shell with only pustules, knobs or nodules ...............................................................................................................................105
97a. (96a)
Shell with undulations, ridges, or plications, but not pustules or nodules on the surface ............................................................98
97b.
Shell with undulations, ridges or plications and pustules on the disc or umbonal area .............................................................103
98a. (97a)
Shell surface with undulations, plications, ridges restricted to the posterior slope ......................................................................99
98b.
Shell surface with undulations, plications, ridges not restricted to the posterior slope ..............................................................101
99a. (98a)
Shell shape round to oval, slablike, compressed, left valve with an accessory dentacle posterior to pseudocardinal teeth plications occur on dorsal wing ...................................................................................Lasmigona [in part] [L. complanata] (Fig. 28X)
99b.
Shell shape more elongate, lacks the accessory dentacle............................................................................................................100
100a. (99b)
Shell inflated with a blue to blue-green nacre color .................................Medionidus [in part] [M. accutissimus, M. mcglameriae,
M. parvulus, M. penicillatus, M. simpsonianus, M. walkeri] (Fig. 28AC)
100b.
Shell inflated, nacre not as above, shell thin, posterior slope steeply angled to moderately angled ..............................Alasmidonta
[in part] [A. triangulata, A. wrightiana] (Fig. 28D)
101a. (98b)
Shell shape rectangular, with a distinct posterior ridge, purple nacre, open and shallow beak cavity ...........................Plectomerus
dombeyanus (Fig. 28AH)
101b.
Shell shape round to rectangular, without a distinct posterior ridge..........................................................................................102
102a. (101b)
Shell thick, compressed, low plications or wrinkles on posterior slope and undulations and ridges on the disk of the shell, nacre
with a central white area and a peripheral purple band, restricted to the Apalachicola and Ochlockonee River basins ..................
..................................................................................................................................................Elliptoideus sloatianus (Fig. 28Q)
102b.
Shell variably inflated, shell shape round to quadrate, heavy pseudocardinal teeth, deep compressed beak cavity, sculpture consists of three to five well-developed ridges or undulations running across the shell from anterior of the umbo to the posterior
ridge, nacre color white to iridescent ...........................................Amblema [in part] [A. plicata, A. neislerii, A. elliotti] (Fig. 28E)
103a. (97b)
Shell shape elongate, with a well-developed diagonal posterior ridge ..........................................Tritogonia verrucosa (Fig. 28AT)
103b.
Shell shape not elongate, lacking well-defined posterior ridge...................................................................................................104
104a (103b).
Shell shape round to quadrate, lacking well-defined posterior ridge, has well developed plications running across the shell and
pustules on the disk or umbonal area .........................................................................................Megalonaias nervosa (Fig. 28AD)
104b.
Plications not well developed and wavy, not straight, plications on posterior slope, two rows of pustules or knobs on umbo........
......................................................................................................................................................Arcidens confragosa (Fig. 28H)
105a. (96b)
Pustules distributed across most of the shell .............................................................................................................................106
105b.
Pustules restricted to a single or double row.............................................................................................................................107
106a. (105a).
Adult shell length 60 mm, sculpture fine, shell oval to elongate oval, pustules and chevrons...................................Quincuncina
[Q. infucata, Q. burkei restricted to the Choctawhatchee River Basin; Q. mitchelli from the Rio Grand, Guadalupe, Colarado
and Brazos rivers in Texas] (Fig. 28AP)
106b.
Adult shell length not restricted to 60 mm, pustules covering most of the shell, nacre color white to pink, not purple, with a
variable number of pustules, from a very few to pustules covering most of the shell, nacre color mostly white, beak cavity deep
and open, not compressed...................................................................................Quadrula [in part] [Q. petrina, Q. houstonensis,
Q. couchiana, Q. aurea, Q. asperata] (Fig. 28AO)
107a. (105b)
Pustules or nodules large, forming a single row down the center of the disk, and limited to about three pustules on each
valve ...............................................................................................................................................Obliquaria reflexa (Fig. 28AE)
107b.
Pustules forming two distinct rows especially on the umbonal area..........................................................................................108
108a. (107b)
Two rows of pustules, with one row on the posterior ridge, shell thick ........................Quadrula [in part] [Q apiculata, has small
pustules in the sulcus between the two rows of pustules; Q. nodulata lacks a sulcus between the rows of pustules and may only
have very few pustules; Q. quadrula, Q. rumphiana] (Fig. 28AO)
108b.
Radiating rows of knobs, shell inflated, thin to relatively thick, umbo high and full, pseudocardinal teeth compressed, not
curved, beak sculpture consists of irregular nodules forming two loops which continue down and across most of the shell in two
radiating rows..............................................................................................................................Arcidens confragosus (Fig. 28H)
414
Robert F. McMahon and Arthur E. Bogan
109a. (95b)
Shell with a well-developed dorsal wing projecting above the hinge line, the wing usually posterior of the umbo [see Fig. 30]
but some species may also have an anterior wing [some older shells such as in Leptodea fragilis may be lacking the dorsal
wing] ........................................................................................................................................................................................110
109b.
Shell without a well-developed dorsal wing ..............................................................................................................................111
110a. (109a)
Pseudocardinal teeth moderately heavy, projecting perpendicular to the axis of the hinge line, purple nacre thin to thick shelled,
may be inflated ...................................................................................................Potamilus [P. inflatus, P. purpuratus] (Fig. 28AL)
110b.
Pseudocardinal teeth compressed, thin and not projecting perpendicular to the axis of the hinge line..............................Leptodea
[L. fragilis, L. amphichaenus] (Fig. 28Z)
111a. (109b)
Shell shape round .....................................................................................................................................................................112
111b.
Shell shape not round ...............................................................................................................................................................117
112a. (111a)
Shell compressed, shell shape round, with lateral teeth, steep posterior slope, shell thick with well-developed teeth, periostracum
light yellow with interrupted rows of green chevrons.......................................................................Ellipsaria lineolata (Fig. 28O)
112b.
Shell not compressed, round to oval .........................................................................................................................................113
113a. (112b)
Umbo central or nearly central in the dorsal margin.................................................................................................................114
113b.
Umbo anterior of the center of the dorsal margin of the shell...................................................................................................116
114a. (113a)
Broad green ray extending around the umbo down only a short distance onto the disk of the shell ................................Quadrula
[in part] [Q. asperata without pustules] (Fig. 28AO)
114b.
No broad green ray ..................................................................................................................................................................115
115a. (114b)
Shell surface smooth, without a broad central sulcus.......................Obovaria [in part] [O. jacksoniana, O. unicolor] (Fig. 28AF)
115b.
Shell with broad, shallow to pronounced sulcus, fine green rays, shallow beak cavity, well developed hinge teeth, female shells
with a swollen, extended or expanded portion of the posterior slope and posterior ventral margin of the shell..............................
...............................................................................Epioblasma [in part] [E. penita, E. metastriata, E. othcaloogensis] (Fig. 28R)
116a. (113b)
Beak cavity deep compressed ........................................................Fusconaia [in part][F. ebena, F. succissa, F. escambia] (Fig. 28S)
116b.
Beak cavity shallow and open, shell shape wedge shaped to triangular or approaching square with or without a central sulcus,
nacre white to deep pink ........................................................................................Pleurobema [in part] [P. marshalli] (Fig. 28AJ)
117a. (111b)
Shell shape not oval..................................................................................................................................................................118
117b.
Shell shape oval ........................................................................................................................................................................130
118a. (117a)
Shell shape triangular, square to rectangular, with sulcus..........................................................................................................119
118b.
Shell shape elongate, tapered posteriorly, two to four times longer than high ...........................................................................123
119a. (118a)
Beak cavity deep.......................................................................................................................................................................120
119b.
Beak cavity shallow or broad and open ....................................................................................................................................121
120a. (119a)
Shell shape rectangular with green rays on the umbo........................................................Fusconaia [in part] [F. cerina] (Fig. 28S)
120b.
Shell shape triangular to broadly triangular ...........................................................Pleurobema [in part] [P. taitianum] (Fig. 28AJ)
121a. (119b)
Posterior slope rounded, not very steep, covered with lines of green chevrons ....................................................Truncilla [in part]
[T. cognata, T. macrodon, T. truncata, T. donaciformis] (Fig. 28AU)
121b.
Posterior ridge very sharp, posterior slope steep .......................................................................................................................122
122a. (121b)
Shell thin, inflated, abruptly truncate posteriorly, posterior slope lighter colored than the rest of the shell .....................................
...........................................................................................................................Alasmidonta [in part] [A. triangulata] (Fig. 28D)
122b.
Shell thin to thick, inflated posterior slope steep shell thick, shell shape rectangular, posterior margin not modified into a marsupial swelling, often tapering posteriorly, nacre varies pale salmon to purple..........................................................Elliptio [in part]
[E. crassidens] (Fig. 28P)
123a. (118b). Thin shelled..............................................................................................................................................................................124
123b.
Thick shelled ............................................................................................................................................................................126
124a. (123a)
Shell compressed, shell shape elongate to almost rectangular, very thin, periostracum rayed, nacre iridescent, white to dull purple, beak cavity shallow, lateral teeth long and curved to straight, pseudocardinal very small, periostracum brown with faint
rays as a juvenile, restricted to the Rio Grande Basin, Texas..............................................................Popenaias popei (Fig. 28AK)
124b.
Shell inflated.............................................................................................................................................................................125
125a. (124b)
Hinge thin, periostracum usually heavily rayed ..................................................................Villosa [in part] [V. nebulosa complex,
V. villosa, V. vibex] (Fig. 28AY)
11. Mollusca: Bivalvia
415
125b.
Hinge not thin, periostracum yellow, plain or rayed ...................................Lampsilis [in part] [L. teres, L. straminea, L. hydiana,
L. bracteata] (Fig. 28W)
126a. (123b)
Shell compressed ......................................................................................................................................................................127
126b.
Shell inflated.............................................................................................................................................................................128
127a. (126a)
Hinge massive, curved, nacre white, periostracum with interrupted green rays....................................... Ptychobranchus [in part]
[P. greeni] (Fig. 28AM)
127b.
Hinge thinner, straight, nacre purple to white, periostracum typically unrayed .....................................................Elliptio [in part]
[E. arctata, E. arca] (Fig. 28P)
128a. (126b)
Rayed as a juvenile becoming dark brown to black as an adult, tapered to a point in middle of posterior margin, nacre white to
white with purple wash in umbo area, Ligumia [L. recta, L. subrostrata] (Fig. 28AB)
128b.
Shell unrayed
129a. (128b)
Adult shell length 50mm, bluntly pointed posterior ventrally, coarse concentric beak sculpture seeming to radiate from a single point, Uniomerus [in part] [U. tetralasmus, U. declivus] (Fig. 28AV)
129b.
Adult shell length 50 mm, bluntly rounded or squared off posteriorly, concentric beak sculpture in form of bars or ridges, angled up posteriorly Toxolasma [in part] [T. parvus, T. texasiensis] (Fig. 28AS)
130a. (117b)
Shell shape oval ........................................................................................................................................................................131
130b.
Shell shape oblong....................................................................................................................................................................133
131a. (130a)
Posterior half of the pseudocardinal tooth divided into several parallel vertical ridges (Fig. 29), thick shelled, inflated, periostracum dark brown to dark olive ......................................................................................................Glebula rotundata (Fig. 28T)
131b.
Pseudocardinal tooth not divided into vertical ridges as above .................................................................................................132
132a. (131b)
Shell shape oval, inflated, rayed, thin-to-thick shell, posterior ridge rounded or sharp, nacre variously white to pink or salmon
..............................................................................................................Lampsilis [in part] [L. ornata, L. binominata] (Fig. 28W)
FIGURE 29 Structure of the posterior pseudocardinal teeth of the
right valve of Gelbula rotundata. Note that the posterior pseudocardinal teeth are deeply divided into parallel, vertical, plicate lamellae,
a tooth arrangement uniquely characteristic of this species. (Redrawn
from Burch, 1975b.)
FIGURE 30 Right shell valve of the unionid, Potamilus alatus,
showing the thin, extensive dorsal projection of the shell posterior to
the umbos forming a “wing” whose presence or absence is a diagnostic characteristic valuable for identification of a number of unionid
species. (Redrawn from Burch, 1975b.)
416
Robert F. McMahon and Arthur E. Bogan
132b.
Shell shape elongate, oval...............................Villosa [in the following two species male shells are oval while female shells have a
truncate posterior margin, V. vanuxemensis umbrans, purple nacre, restricted to the upper Coosa River Basin, V. lienosa, variously colored nacre] (Fig. 28AY)
133a. (130b)
Shell shape oblong, inflated to globose, unrayed........................Potamilus [in part] [P. amphichaenus, P. purpuratus] (Fig. 28AL)
133b.
Shell shape oblong, inflated, but not globose ............................................................................................................................134
134a. (133b)
Shell shape elongate oval, shell relatively thin to thick, compressed, pseudocardinal teeth compressed, thin, lateral teeth long,
beak cavity shallow, periostracum tan to brown, with dark brown rays posteriorly, restricted to the Rio Grande Basin in
Texas................................................................................................Disconaias [in part] [D. salinasensis, D. conchos] (Fig. 28M)
134b.
Shell shape oval, moderately thick shell, inflated, beak cavity relatively deep, periostracum dull/shiny, red-brown to brown,
juveniles with faint rays, Brazos River to Rio Grande Basin, Texas ..........................................Cyrtonaias tampicoensis (Fig. 28L)
ACKNOWLEDGMENTS
Alan P. Covich and Caryn C. Vaugh critically reviewed the chapter and made many valuable suggestions for its improvement. Thomas H. Dietz, Daniel
Hornbach, Carl M. Way, and Harold Silverman provided the scanning and transmission electron micrographs of bivalve gill ciliation presented in Figure 21.
Paul W. Parmalee, Caryn C. Vaughn, Dan Spooner, Valeria Rice and Jeff Garner kindly reviewed and commented on the unionoidean key. Jonathon A. Raine
and Chad Rubins provided technical support with the
preparation of the illustrations used in Figure 28.
Cindy Bogan provided suggestions and encouragement
during the drafting of the unionoidean key. Mrs. W. T.
Edmondson kindly granted her permission for us to use
the figures from her late husband’s second edition of
Ward and Wipple (Edmondson, 1959).
LITERATURE CITED
Aarset, A. V. 1982. Freezing tolerance in intertidal invertebrates (a
review). Comparative Biochemistry and Physiology A73:
571 – 580.
Adam, M. E. 1986. The Nile bivalves: how do they avoid silting during the flood? Journal of Molluscan Studies 52:248 – 252.
Ahlstedt, S. A. 1983. The Mollusca of the Elk River in Tennessee and
Alabama. American Malacological Bulletin 1:43 – 50.
Aldridge, D. W., McMahon, R. F. 1978. Growth, fecundity, and
bioenergetics in a natural population of the Asiatic freshwater
clam, Corbicula manilensis Philippi, from north central Texas.
Journal of Molluscan Studies 44:49 – 70.
Aldridge, D. W., Payne, B. S., Miller, A. C. 1987. The effect of intermittent exposure to suspended solids and turbulence on three
species of freshwater mussels. Environmental Pollution
45:17 – 28.
Alexander, J. E., Jr., Thorp, J. H., Fell, R. D. 1994. Turbidity and
temperature effects on oxygen consumption in the zebra mussel
(Dreissena polymorpha). Canadian Journal of Fisheries and
Aquatic Science 51:179 – 184.
Alimov, A. F. 1975. The rate of metabolism of freshwater bivalve
molluscs. Soviet Journal of Ecology 6:6 – 13.
Allen, A. J. 1985. Recent Bivalva: their form and evolution. in: Trueman, E. R., Clarke, M. R., Eds. The Mollusca, Vol. 10, Evolution. Academic Press, New York, pp. 337 – 403.
Allen, Y. C., Thompson, B. A., Ramcharan, C. W. 1999. Growth and
mortality rates of the zebra mussel, Dreissena polymorpha, in
the lower Mississippi River. Canadian Journal of Fisheries and
Aquatic Science 56:748 – 759.
Anderson, R. M., Layzer, J. B., Gordon, M. E. 1991. Recent catastrophic decline of mussels (Bivalvia: Unionidae) in the Little
South Fork Cumberland River, Kentucky. Brimleyana 17:1 – 8.
Annis, C. G., Belanger, T. V. 1986. Corbicula manilensis, potential
bio–indicator of lead and copper pollution. Florida Scientist
49:30.
Araujo, R., Remón, J. M., Moreno, D., Ramos, M. A. 1995. Relaxing techniques for freshwater molluscs: trials for evaluation of
different methods. Malacologia 36:29 – 41.
Arnott, D. L., Vanni, M. J. 1996. Nitrogen and phosphorous recycling by the zebra mussel (Dreissena polymorpha) in the Western Basin of Lake Erie. Canadian Journal of Fisheries and
Aquatic Science 53:646 – 659.
Arter, H. E. 1989. Effect of eutrophication on species composition
and growth of freshwater mussels (Mollusca, Unionidae) in Lake
Hallwil (Aargau, Switzerland). Aquatic Sciences 51:87 – 99.
Baily, R. C., 1989. Habitat selection by a freshwater mussel: an experimental test. Malacologia 31:205 – 210.
Baily, R. C., Green, R. H. 1989. Spatial and temporal variation in a
population of freshwater mussels in Shell Lake, N.W.T. Canadian Journal of Fisheries and Aquatic Science 46:1392 – 1395.
Bauer, G. 1983. Age structure, age specific mortality rates and population trend of the freshwater pearl mussel (Margaritifera margaritifera) in north Bavaria. Archiv für Hydrobiologie 98: 523 – 532.
Bauer, G. 1994. The adaptive value of offspring size among freshwater mussels (Bivalvia; Unionidae). Journal of Animal Ecology
63:933 – 944.
Belanger, S. E. 1991. The effect of dissolved oxygen, sediment and
sewage treatment plant discharges upon growth, survival and
density of Asiatic clams. Hydrobiologia 218:113 – 126.
Belanger, S. E., Farris, J. L., Cherry, D. S., Cairns, J., Jr., 1985. Sediment preference of the freshwater Asiatic clam, Corbicula fluminea. Nautilus 99:66 – 73.
Belanger, S. E., Cherry, D. S., Cairns, J., Jr. 1986a. Uptake of
chrysotile asbestos fibers alters growth and reproduction of Asiatic clams. Canadian Journal of Fisheries and Aquatic Sciences
43:43 – 52.
Belanger, S. E., Farris, J. L., Cherry, D. S., Cairns, J., Jr. 1986b.
Growth of Asiatic clams (Corbicula sp.) during and after longterm zinc exposure in field-located and laboratory artificial
streams. Archives of Environmental Contamination and Toxicology 15:427 – 434.
Belanger, S. E., Cherry, D. S., Cairns, J., Jr., McGuire, M. J. 1987.
Using Asiatic clams as a biomonitor for chrysotile asbestos in
public water supplies. Journal of the American Water Works
Association 79:69 – 74.
11. Mollusca: Bivalvia
Beninger, P. G., Lynn, J. W., Dietz, T. H., Silverman, H. 1997. Mucociliary transport in living tissue: two-layer model confirmed in
the mussel, Mytilus edulis L. Biological Bulletin (Woods Hole,
Mass.) 193:4 – 7.
Bisbee, G. D. 1984. Ingestion of phytoplankton by two species of
freshwater mussels, the black sandshell, Ligumia recta, and the
three ridger, Amblema plicata, from the Wisconsin River in
Oneida County, Wisconsin. Bios 58:219 – 225.
Bishop, S. H., Ellis, L. L., Burcham, J. M. 1983. Amino acid metabolism in molluscs. in: Hochachka, P. W., Ed. The Mollusca. Vol.
1: Metabolic biochemistry and molecular biomechanics. Academic Press, New York, pp. 243 – 327.
Bitterman, A. M., Hunter, R. D., Haas, R. C. 1994. Allometry of
shell growth of caged and uncaged mussels (Dreissena polymorpha) in Lake St. Clair. American Malacological Bulletin
11:41 – 49.
Blalock, H. N., Sickel, J. B. 1996. Changes in mussel (Bivalvia:
Unionidae) fauna within the Kentucky portion of Lake Barkley
since impoundment of the lower Cumberland River. American
Malacological Bulletin 13:111 – 116.
Blay, J., Jr. 1990. Fluctuations in some hydrological factors and the
condition index of Aspatharia sinuata (Bivalvia, Unionacea) in a
small Nigerian Reservoir. Archiv für Hydrobiologie
117:357 – 363.
Blye, R. W., Ettinger, W. S., Nebane, W. N. 1985. Status of Asiatic
clam (Corbicula fluminea) in south eastern Pennsylvania-1984.
Proceedings of the Pennsylvania Academy of Science 59:74.
Bogan, A. E. 1990. Stability of recent unionid (Mollusca: Bivalvia)
communities over the past 6000 years. in: Miller, W., III, Ed.
Paleocommunity temporal dynamics: the long-term development of multispecies assemblages. The Paleontological Society,
Special Publication No. 5, Washington, DC, pp. 112 – 136.
Bogan, A. E. 1993. Freshwater bivalve extinctions: search for a
cause. American Zologist 33:599 – 609.
Bogan, A. E. 1997. The silent extinction. American Paleontologist
15:2 – 4.
Bogan, A. E. 1998. Freshwater molluscan conservation in North
America: problems and practices. in: Seddon, M. B., Holmes, A.
M., Eds. Molluscan conservation: a strategy for the 21st Century. I. J. Killeen, Journal of Conchology, Special Publication
No. 2, pp. 223 – 230.
Bogan, A. E., Woodward, F. R. 1992. A review of the higher classification of Unionoida (Mollusca: Bivalvia). Unitas Malacologica
Abstracts of the Eleventh International Malacological Congress,
Siena, Italy, pp. 17 – 18.
Bonaventura, C., Bonaventura, J. 1983. Respiratory pigments: structure and function. in: Hochachka, P. W., Ed. The Mollusca. Vol.
2: Environmental chemistry and physiology. Academic Press,
New York, pp. 1 – 50.
Borcherding, J. 1992. Another early warning system for the detection
of toxic discharges in aquatic environments based on valve
movements of the freshwater mussel, Dreissena polymorpha.
Limnologie Aktuell 4:127 – 146.
Borcherding, J., Volpers, M. 1994. The “Dreissena-monitor” — first
results on the application of this biological early warning system in the continuous monitoring of water quality. Water Science Technology 29:199 – 201.
Botts, P. S, Patterson, B. A., Schloesser, D. W. 1996. Zebra mussel effects on benthic invertebrates: physical or biotic? Journal of the
North American Benthological Society 15:179 – 184.
Britton, J. C., Morton, B. 1986. Polymorphism in Corbicula Fluminea (Bivalvia: Corbiculidae) from North America. Malacological Review 19:1 – 43.
Brown, K. M., Curole, J. P. 1997. Longitudinal changes in the mussels of the Amite River: endangered species, effects of gravel
417
mining and shell morphology. in: Cummings, K. S., Buchanan,
A. C., Mayer, C. A., Naimo, T. J., Eds. Conservation and management of freshwater mussels II: Initiatives for the future, Proceedings of a Symposium, 1995, Upper Mississippi River Conservation Committee, Rock Island, IL, pp. 236 – 246.
Buddensiek, V., Engel, H., Fleischauer-Rössing, S., Wächtler, K.
1993. Studies on the chemistry of interstitial water taken from
defined horizons in the fine sediments of bivalve habitats in
several northern German lowland waters. II: Microhabitats of
Margaritifera margaritifera L., Unio crassus (Philipsson), and
Unio tumidus Philipsson. Archive feur Hydrobiologie
127:151 – 166.
Burch, J. B. 1975a. Freshwater sphaeriacean clams (Mollusca: Pelecypoda) of North America. Malacological Publications, Hamburg,
MI.
Burch, J. B. 1975b. Freshwater unionacean clams (Mollusca: Pelecypoda) of North America. Malacological Publications, Hamburg,
MI.
Burky, A. J. 1983. Physiological ecology of freshwater bivalves. in:
Russell-Hunter, W. D., Ed. The Mollusca. Vol. 6 : Ecology. Academic Press, New York, pp. 281 – 327.
Burky, A. J., Burky, K. A. 1976. Seasonal respiratory variation and
acclimation in the pea clam, Pisidium walkeri Sterki. Comparative Biochemistry and Physiology. A55:109 – 114.
Burky, A. J., Hornbach, D. J., Way, C. M. 1981. Growth of Pisidium
casertanum (Poli) in west central Ohio. Ohio Journal of Science
81:41 – 44.
Burky, A. J., Benjamin, R. B., Conover, D. G., Detrick, J. R. 1985a.
Seasonal responses of filtration rates to temperature, oxygen
availability, and particle concentration of the freshwater clam
Musculium partumeium (Say). American Malacological Bulletin
3:201 – 212.
Burky, A. J., Hornbach, D. J., Way, C. M. 1985b. Comparative
bioenergetics of permanent and temporary pond populations of
the freshwater clam, Musculium partumeium (Say). Hydrobiologia 126:35 – 48.
Burnett, L. E. 1997. The challenges of living in hypoxic and hypercapnic aquatic environments. American Zoologist 37:633 – 640.
Burton, R. F. 1983. Ionic regulation and water balance. in: Saleuddin, A. S. M., Wilbur, K. M., Ed. The Mollusca. Vol. 5: Physiology, Part 2. Academic Press, Orlando, FL, pp. 292 – 352.
Buttner, J. K. 1986. Corbicula as a biological filter and polyculture
organism in catfish rearing ponds. Progessive Fish-Culturist
48:136 – 139.
Butts, T. A., Sparks, R. E. 1982. Sediment oxygen demand-fingernail
clam relationship in the Mississippi River Keokuk Pool. Transactions of the Illinois Academy of Sciences 75:29 – 39.
Byrne, R. A., Dietz, T. H. 1997. Ion transport and acid-base balance
in freshwater bivalves. Journal of Experimental Biology
200:457 – 465.
Byrne, R. A., McMahon, B. R. 1991. Acid – base and ionic regulation, during and following emersion, in the freshwater bivalve,
Anodonta grandis simpsoniana (Bivalvia: Unionidae). Biological
Bulletin (Woods Hole, Mass.) 181:289 – 297.
Byrne, R. A., McMahon, R. F. 1994. Behavioral and physiological responses to emersion in freshwater bivalves. American Zoologist
34:194 – 204.
Byrne, R. A., McMahon, R. F., Dietz, T. H. 1988. Temperature and
relative humidity effects on aerial exposure tolerance in the
freshwater bivalve Corbicula fluminea. Biological Bulletin
(Woods Hole, Mass.) 175:253 – 260.
Byrne, R. A., Gnaiger, E., McMahon, R. F., Dietz, T. H. 1990. Behavioral and metabolic responses to emersion and subsequent reimmersion in the freshwater bivalve, Corbicula fluminea. Biological Bulletin (Woods Hole, Mass.) 178:251 – 259.
418
Robert F. McMahon and Arthur E. Bogan
Byrne, R. A., Shipman, B. N., Smatresk, N. J., Dietz, T. H., McMahon, R. F. 1991a. Acid – base balance during emergence in the
freshwater bivalve Corbicula fluminea. Physiological Zoology
64:748 – 766.
Byrne, R. A., Dietz, T. H., McMahon, R. F. 1991b. Ammonia dynamics during and after prolonged emersion in the freshwater
clam Corbicula fluminea (Müller) (Bivalvia: Corbiculacea).
Canadian Journal of Zoology 69:676 – 680.
Cáceres, C. E. 1997. Dormancy in invertebrates. Invertebrate Biology
116:371 – 383.
Cairns, J., Jr., Cherry, D. S. 1983. A site-specific field and laboratory
evaluation of fish and Asiatic clam population responses to coal
fired power plant discharges. Water Science and Technology
15:31 – 58.
Calow, P. 1983. Life-cycle patterns and evolution. in: Russell-Hunter,
W. D., Ed. The Mollusca. Vol. 6: Ecology. Academic Press, New
York, pp. 649 – 678.
Clarke, A. H. 1973. The freshwater molluscs of the Canadian interior basin. Malacologia 13:1 – 509.
Clarke, A. H. 1983. The distribution and relative abundance of
Lithasia pinguis (Lea), Pleurobema plenum (Lea), Villosa trabalis (Conrad), and Epioblasma sampsoi (Lea). American Malacological Bulletin 1:27 – 30.
Clarke, M., McMahon, R. F., Miller, A. C., Payne, B. S. 1993. Tissue
freezing points and time for complete mortality on exposure to
freezing air temperatures in the zebra mussel (Dreissena polymorpha) with special reference to dewatering during freezing
conditions as a mitigation strategy, in: Tsou, J. L., Muussalli, Y.
G., Eds. Proceedings: Third International Zebra Mussel Conference 1993, EPRI TR-102077, Electric Power Research Institute,
Palo Alto, CA, pp. 1 – 145.
Clarke, M., McMahon, R. F. 1996a. Comparison of byssal attachment in dreissenid and mytilid mussels: mechanisms, morphometry, secretion, biochemistry, mechanics and environmental influences. Malacological Review 29:1 – 16.
Clarke, M., McMahon, R. F. 1996b. Effects of hypoxia and low-frequency agitation on byssogenesis in the freshwater mussel
Dreissena polymorpha. Biological Bulletin (Woods Hole, Mass.)
191:413 – 420.
Claudi, R., Mackie, G. L. 1993. Practical manual for zebra mussel
monitoring and control. Lewis Publishers, Boca Raton, FL.
Cleland, J. D., McMahon, R. F., Elick, G. 1986. Physiological differences between two morphotypes of the Asian clam. Corbicula.
American Zoologist 26:103A.
Clench, W. J. 1959. Mollusca. in: Edmondson, W. T., Ed. Freshwater
biology. 2nd ed., Wiley, New York, pp. 1117 – 1160.
Clench, W. J., Turner, R. D. 1956. Freshwater mollusks of Alabama,
Georgia, and Florida from the Escambia to the Suwannee River.
Bulletin of the Florida State Museum, Biological Sciences
1:97 – 239.
Cohen, R. R. H., Dressler, P. V., Phillips, E. P. J., Cory, R. L. 1984.
The effect of the Asiatic clam, Corbicula fluminea, on phytoplankton of the Potomac River, Maryland. Limnology and
Oceanography 29:170 – 180.
Collins, T. W. 1967. Oxygen-uptake, shell morphology and dessication of the fingernail clam, Sphaerium occidentale Prime. Ph.D.
Thesis, University of Minnesota, Minneapolis.
Coney, C. C. 1993. An empirical evaluation of various techniques for
anesthetization and tissue fixation of freshwater Unionoida
(Mollusca: Bivalvia), with a brief history of experimentation in
molluscan anesthetization. Veliger 36:413 – 424.
Conn, D. B., Ricciardi, A., Babapulle, M. N., Klien, K. A., Rosen, D.
A. 1996. Chaetogaster limnaei as a parasite of the zebra mussel
Dreissena polymorpha, and the quagga mussel Dreissena bugensis (Mollusca: Bivalvia). Parasitology Research 82:1 – 7.
Cotner, J. B., Gardner, W. S., Johnson, J. R., Sada, R. H., Cavaletto,
J. F., Heath, R. T. 1995. Effects of zebra mussels (Dreissena
polymorpha) on bacterioplankton: evidence for both size-selective consumption and growth stimulation. Journal of Great
Lakes Research 21:517 – 528.
Counts, C. L., III, Handwerker, T. S., Jesien, R. V. 1991. The Naiades
(Bivalvia: Unionoidea) of the Delmarva Peninsula. American
Malacological Bulletin 9:27 – 37
Covich, A. P., Dye, L. L., Mattice, J. S. 1981. Crayfish predation on
Corbicula under laboratory conditions. American Midland Naturalist 105:181 – 188.
Cunjak, R. A., McGladdery, S. E. 1991. The parasite – host relationship of glochidia (Mollusca: Margaritiferidae) on the gills of
young of the year Atlantic salmon (Salmo salar). Canadian
Journal of Zoology 69:353 – 358.
Darwin, C. 1982. On the dispersal of freshwater bivalves. Nature 25:
529.
Das, V. M. M., Venkatachari, S. A. T. 1984. Influence of varying
oxygen tension on the oxygen consumption of the freshwater
mussel Lamellidens marginalis (Lamarck) and its relation to
body size. Veliger 26:305 – 310.
Davis, D. S., Gilhen, J. 1982. An observation of the transportation of
pea clams, Pisidium adamsi, by bluespotted salamanders, Ambystoma laterale. The Canadian Naturalist 96:213 – 215.
Davis, W. L., Jones, R. G., Knight, J. P., Hagler, H. K. 1982. An electron microscopic histochemical and X-ray microprobe study of
spherites in a mussel. Tissue and Cell 14:61 – 67.
Deaton, L. E. 1982. Tissue (NA K)-activated adenosinetriphosphatase activities in freshwater and brackish water bivalve molluscs. Marine Biology Letters 3:107 – 112.
Deaton, L. E., Greenberg, M. J. 1991. The adaptation of bivalve
molluscs to oligohaline and fresh waters: phylogenetic and
physiological aspects. Malacological Review 24:1 – 18.
Denson, D. R., Wang, S. Y. 1994. Morphological differences between
zebra and quagga mussel spermatozoa. American Malacological
Bulletin 11:79 – 81.
de Zwaan, A. 1983. Carbohydrate catabolism in bivalves. in:
Hochachka, P. W., Ed. The Mollusca. Vol. 1: Metabolic biochemistry and molecular biomechanics. Academic Press, New
York, pp. 137 – 175.
Dietz, T. H. 1974. Body fluid composition and aerial oxygen consumption in the freshwater mussel, Ligumia subrostrata (Say):
effects of dehydration and anoxic stress. Biological Bulletin
(Woods Hole, Mass.) 147:560 – 572.
Dietz, T. H. 1985. Ionic regulation in freshwater mussels: a brief review. American Malacological Bulletin 3:233 – 242.
Dietz, T. H., Byrne, R. A. 1997. Effects of salinity on solute clearance
from the freshwater bivalve, Dreissena polymorpha Pallas. Experimental Biology Online 2:11 – 20.
Dietz, T. H., Byrne, R. A. 1999. Measurement of sulfate uptake and
loss in the freshwater bivalve Dreissena polymorpha using semimicroassay. Canadian Journal of Zoology 77:331 – 336.
Dietz, T. H., Scheide, J. I., Saintsing, D. G. 1982. Monoamine
transmitters and cAMP stimulation of Na transport in
freshwater mussels. Canadian Journal of Zoology 60:
1408 – 1411.
Dietz, T. H., Wilson, J. M., Silverman, H. 1992. Changes in
monoamine transmitter concentration in freshwater mussel tissues. Journal of Experimental Zoology 261:355 – 358.
Dietz, T. H., Lessard, D., Silverman, H., Lynn, J. W. 1994. Osmoregulation in Dreissena polymorpha: the importance of Na, Cl, K,
and particularly Mg. Biological Bulletin (Woods Hole, Mass.)
187:76 – 83.
Dietz, T. H., Byrne, R. A., Lynn, J. W., Silverman, H. 1995. Paracellular solute uptake by the freshwater zebra mussel, Dreissena
11. Mollusca: Bivalvia
polymorpha. American Journal of Physiology 269 (Regulatory
and Integrative Comparative Physiology 38):R300 – R307.
Dietz, T. H., Wilcox, S. J., Byrne, R. A., Lynn, J. W., Silverman, H.
1996a. Osmotic and ionic regulation of North American zebra
mussels (Dreissena polymorpha). American Zoologist 36:
364 – 372.
Dietz, T. H., Wilcox, S. J., Byrne, R. A., Silverman, H. 1996b. Effects
of hyperosmotic challenge on the freshwater bivalve Dreissena
polymorpha: Importance of K. Canadian Journal of Zoology
75:697 – 705.
Dietz, T. H., Neufeld, D. H., Silverman, H., Wright, S. H. 1998. Cellular volume regulation in freshwater bivalves. Journal of Comparative Physiology B 168:87 – 95.
Di Maio, J., Corkum, L. D. 1995. Relationship between the spatial
distribution of freshwater mussels (Bivalvia: Unionidae) and the
hydrological variability of rivers. Canadian Journal of Zoology
73:663 – 671
Di Maio, J., Corkum, L. D. 1997. Patterns of orientation in unionids
as a function of rivers with differing hydrological variability.
Journal of Molluscan Studies 63:531 – 539.
Dimock, R.V., Jr., Wright, A. H. 1993. Sensitivity of juvenile freshwater mussels to hypoxic, thermal and acid stress. The Journal
of the Elisha Mitchell Scientific Society 109:183 – 192.
Doherty, F. G., Cherry, D. S., Cairns, J., Jr. 1987. Valve closure responses of the Asiatic clam Corbicula fluminea exposed to cadmium and zinc. Hydrobiologia 153:159 – 167.
Downing, W. L., Shostell, J., Downing, J. A. 1992. Non–annual external
annuli in the freshwater mussels Anodonta grandis grandis and
Lampsilis radiata siliquiodea. Freshwater Biology 28:309– 317.
Downing, J. A., Rochon, Y., Pérusse, M., Harvey, H. 1993. Spatial
aggregation, body size, and reproductive success in the freshwater mussel, Elliptio complanata. Journal of the North American
Benthological Society 12:148 – 156.
Dreier, H. 1977. Study of Corbicula in Lake Sangchris. in: The annual report for fiscal year 1976, Lake Sangchris project, section
7. Illinois Natural History Survey, Urbana, IL, pp. 7.1 – 7.52.
Dreier, H., Tranquilli, J. A. 1981. Reproduction, growth, distribution, and abundance of Corbicula in an Illinois cooling lake.
Illinois Natural History Survey Bulletin 32:378 – 393.
Duncan, R. E., Thiel, P. A. 1983. A survey of the mussel densities in
pool 10 of the upper Mississippi River. Technical Bullletin No.
139. Wisconsin Department of Natural Resources, Madison. 14
pp.
Dyduch-Falniowska, A. 1982. Oscillations in density and diversity of
Pisidium communities in two biotopes in southern Poland. Hydrobiological Bulletin 16:123 – 132.
Edmondson, W. T. (Ed.). 1959. Freshwater biology (second edition).
Wiley, New York.
Effler, S. W., Siegfried, C. 1994. Zebra mussel (Dreissena polymorpha)
populations in the Seneca River, New York: impact on oxygen resources. Environmental Science and Technology 28:2216 – 2221.
Elder, J. F., Mattraw, H. C., Jr. 1984. Accumulation of trace elements,
pesticides, and polychlorinted biphenyls in sediments and the clam
Corbicula manilensis of the Apalachicola River, Florida. Archives
of Environmental Contamination and Toxicology 13:453 – 469.
Ellis, A. E. 1978. British freshwater bivalve Mollusca. Keys and notes
for identification of the species. Academic Press, New York.
Eng, L. L. 1979. Population dynamics of the Asiatic clam, Corbicula
fluminea (Müller), in the concrete-lined Delta – Mendota Canal
of central California. in: Britton, J. C., Ed. Proceedings, first international Corbicula symposium. Texas Christian University
Research Foundation, Fort Worth, TX, pp. 39 – 68.
Englund, V., Heino, M. 1994a. Valve movement of Anodonta
anatina and Unio tumidus (Bivalvia, Unionidae) in a eutrophic
lake. Annales Zoologici. Fennici 31:257 – 262.
419
Englund, V. P. M., Heino, M. P. 1994b. A new method for the identification marking of bivalves in still water. Malacological Review
27: 111 – 112.
Ferris, J. L., van Hassel, J. H., Belanger, S. E., Cherry, D. S., Cairns,
J., Jr. 1988. Application of cellulolytic activity of Asiatic clams
(Corbicula sp.) to instream monitoring of power plant effluents.
Environmental Toxicology and Chemistry 7:701 – 713.
Fisher, S. W., Gossiaux, D. G., Bruner, K. A., Landrum, P. F. 1993.
Investigation of the toxicokinetics of hydrophobic contaminants
in zebra mussel. in: Nalepa, T. F., Schloesser, D., Eds. Zebra
mussels: biology, impacts, and control. Lewis Publishers, Boca
Raton, FL, pp. 465 – 488.
Florkin, M., Bricteux-Gregoire, S. 1972. Nitrogen metabolism in molluscs. in: Florkin, M., Scheer, B. T., Eds. Chemical zoology, Vol.
VII, Mollusca. Academic Press, San Diego, CA, pp. 301 – 348.
Focarelli, R., Rosa, D., Rosati, F. 1990. Differentiation of the
vitelline coat and the polarized site of sperm entrance in the egg
of Unio elongatulus (Mollusca, Bivalvia). Journal of Experimental Zoology 254:88 – 96.
Foe, C., Knight, A. 1985. The effect of phytoplankton and suspended
sediment on the growth of Corbicula fluminea (Bivalvia). Hydrobiologia 127:105 – 115.
Foe, C., Knight, A. 1986a. A thermal energy budget for juvenile Corbicula fluminea. American Malacological Bulletin, Special ed.
No. 2:143 – 150.
Foe, C., Knight, A. 1986b. Growth of Corbicula fluminea (Bivalvia)
fed on artificial and algal diets. Hydrobiologia 133:155 – 164.
Foe, C., Knight, A. 1986c. A method for evaluating the sublethal impact of stress employing Corbicula fluminea. American Malacological Bulletin, Special ed. No. 2:133 – 142.
Foe, C., Knight, A. 1987. Assessment of the biological impact of
point source discharges employing Asiatic clam. Archives of Environmental Contamination and Toxicology 16:39 – 51.
French, J. R. P., III, Love, J. G. 1995. Size limitation on zebra mussels
consumed by freshwater drum may preclude the effectiveness of
drum as a biological controller. Journal of Freshwater Ecology
10:379 – 383.
Fritz, L. W., Lutz, R. A. 1986. Environmental perturbations reflected
in internal shell growth patters of Corbicula fluminea (Mollusca: Bivalvia). Veliger 28:401 – 417.
Fukuhara, S, Nagata, Y. 1988. Frequency of incubation of Anodonta
woodiana in small pond. Venus 47:271 – 277.
Fuller, S. L. H. 1974. Clam and mussels (Mollusca: Bivalvia). in:
Hart, C. W., Jr., Fuller, S. L. H., Eds. Pollution ecology of freshwater invertebrates. Academic Press, New York, pp. 215 – 273.
Gainey, L. F., Jr., Greenberg, M. J. 1977. Physiological basis of the
species abundance-salinity relationship in molluscs: a speculation. Marine Biology (Berlin) 40:41 – 49.
Gardiner, D. B., Silverman, H., Dietz, T. H. 1991. Musculature associated with the water canals in freshwater mussels and response
to monoamines in vitro. Biological Bulletin (Woods Hole,
Mass.) 180:453 – 465
Golightly, C. G., Jr. 1982. Movement and growth of Unionidae
(Mollusca: Bivalvia) in the Little Brazos River, Texas. Ph.D.
Thesis, Texas A&M University, College Station, Tx.
Graf, D. L. 1997. Sympatric speciation of freshwater mussels (Bivalvia:
Unionoidea): a model. American Malacological Bulletin 14:35–40.
Graney, R. L., Jr., Cherry, D. S., Cairns, J., Jr. 1984. The influence of
substrate, pH, diet and temperature upon cadmium accumulation in the Asiatic clam (Corbicula fluminea) in laboratory
streams. Water Research 18:833 – 842.
Grapentine, L. C. 1992. Growth rates of unionid bivalves upstream
and downstream of industrial outfalls contaminated with trace
metals. Canadian Technical Report of Fisheries and Aquatic Sciences 1863:141 – 143.
420
Robert F. McMahon and Arthur E. Bogan
Graves, S. Y., Dietz, T. H. 1980. Diurnal rhythms of sodium transport in the freshwater mussel. Canadian Journal of Zoology
58:1626 – 1630.
Graves, S. Y., Dietz, T. H. 1982. Cyclic AMP stimulation and
prostaglandin inhibition of Na transport in freshwater mussels.
Comparative Biochemistry and Physiology A 71:65 – 70.
Gordon, M. E., Smith, D. G. 1990. Autumnal reproduction in Cumberlandia monodonta (Unionoidea: Margaritiferidae). Transactions of the American Microscopical Society 109:407 – 411.
Gunning, G. E., Suttkus, R. D. 1985. Reclamation of the Pearl River.
A perspective of unpolluted versus polluted waters. Fisheries
10:14 – 16.
Haag, W. R., Warren, M. L., Jr. 1998. Role of ecological factors and
reproductive strategies in structuring freshwater mussel communities. Canadian Journal of Fisheries and Aquatic Science
55:297 – 306.
Haag, W. R., Warren, M. L., Jr. 1999. Mantle displays of freshwater
mussels elicit attacks from fish. Freshwater Biology 42:35 – 40.
Haag, W. R., Butler, R. S., Hartfield, P. D. 1995. An extraordinary
reproductive strategy in freshwater bivalves: prey mimicry to facilitate larval dispersal. Freshwater Biology 34:471 – 476.
Haag, W. R., Warren, M. L., Jr., Shillingsford, M. 1999. Host fishes
and host-attracting behavior of Lampsilis altilis and Villosa
vibex (Bivalvia: Unioniodae). American Midland Naturalist
141:149 – 157.
Haas, F. 1969. Superfamily Unionacea Fleming, 1828. in: Moore, R.
C., Ed. Treatise on invertebrate paleontology. Part N: Mollusca.
The Geological Society of America, Boulder, CO, pp. 411 – 467.
Hakenkamp, C. C., Palmer, M. A. 1999. Introduced bivalves in freshwater ecosystems: the impact of Corbicula on organic matter dynamics in a sandy bottom stream. Oecologia 119:445 – 451.
Hameed, P. S., Raj, A. I. M. 1990. Freshwater mussel, Lamellidens
marginalis (Lamark) (Mollusca: Bivalvia: Unionidae) as an indicator of river pollution. Chemistry and Ecology 4:57 – 64.
Hanson, J. M., McKay, W. C., Prepas, E. E. 1988. The effects of water depth and density on the growth of a unionid clam. Freshwater Biology 19:345 – 355.
Hanson, J. M., MacKay, W. C., Prepas, E. E. 1989. Effect of size-selective predation by muskrats (Ondatra zebithicus) on a population of unionid clams (Anodonta gradis simpsoniana). Journal
of Animal Ecology 58:15 – 28.
Harmon, J. L., Joy, J. E. 1990. Growth rates of the freshwater
mussel, Anodonta imbecillis Say 1829, in five West Virginia
wildlife station ponds. American Midland Naturalist
124:372 – 378.
Hartfield, P. 1993. Headcuts and their effects on freshwater mussels.
in: Cummings, K. S., Buchanan, A. C., Koch, L. M., Eds. Conservation and Management of Freshwater Mussels, Proceedings
of a Symposium, 1992, Upper Mississippi River Conservation
Committee, Rock Island, IL, pp. 131 – 141.
Hartfield, P., Rummel, R. G. 1985. Freshwater mussels (Unionidae)
of the Big Black River, Mississippi. Nautilus 99:116 – 119.
Hartley, D. M., Johnston, J. B. 1983. Use of the freshwater clam
Corbicula manilensis as a monitor for organochlorine pesticides. Bulletin of Environmental Contamination and Toxicology
31:33 – 40.
Haukioja, E., Hakala, T. 1978. Life-history evolution in Anodonta
piscinails (Mollusca, Pelecypoda). Oecologia 35:253 – 266.
Havlik, M. E. 1983. Naiad mollusk populations (Bivalvia: Unionidae) in Pools 7 and 8 of the Mississippi River near La Crosse,
Wisconsin. American Malacological Bulletin 1:51 – 60.
Hawkins, A. J. S., Widdows, J., Bayne, B. L. 1989. The relevance of
whole-body protein metabolism to measured costs of maintenance and growth in Mytilus edulis. Physiological Zoology
62:745 – 763.
Heard, W. H. 1977. Reproduction of fingernail clams (Sphaeriidae:
Sphaerium and Musculium). Malacologia 16:421 – 455.
Hedegaard, C., Wenk, H-R. 1998. Microstructure and texture
patterns of mollusc shells. Journal of Molluscan Studies
64:133 – 136.
Hemelraad, J., Herwig, H. J., Holwerda, D. A., Zandee, D. I. 1985. Accumulation, distribution and localization of cadmium in the freshwater clam Anodonta sp. Marine Environmental Research 17:196.
Heming, T. A., Vinogradov, G. A., Klerman, A. K., Komov, V. T.
1988. Acid – base regulation in the freshwater pearl mussel Margaritifera margaritifera: effect of emersion and low water pH.
Journal of Experimental Biology 137:501 – 511.
Henery, R. P., Saintsing, D. G. 1983. Carbonic anhydrase activity
and ion regulation in three species of osmoregulating bivalve
molluscs. Physiological Zoology 56:274 – 280.
Hernandez, M. R., McMahon, R. F., Dietz, T. H. 1995. Investigation
of geographic variation in the thermal tolerance of zebra mussels, Dreissena polymorpha. in: Proceedings of the fifth zebra
mussel and other aquatic nuisance organisms conference 1995.
Ontario Hydro, Ont., Canada, pp. 195 – 209.
Herwig, H. J., Hemelraad, J., Howerda, D. A., Zandee, D. I. 1985.
Cytochemical localization of cadmium and tin in bivalves. Marine Environmental Research 17:196 – 197.
Higgins, F. 1858. A catalogue of the shell-bearing species, inhabiting
the vicinity of Columbus, Ohio, with some remarks thereon.
Twelfth Annual Report, Ohio Sate Board of Agriculture for
1857:548 – 555.
Hillis, D. M., Mayden, R. L. 1985. Spread of the Asiatic clam, Corbicula (Bivalvia: Corbiculacea), into the new world tropics.
Southwestern Naturalist 30:454 – 456.
Hillis, D. M., Patton, J. C. 1982. Morphological and electrophoretic
evidence for two species of Corbicula (Bivalvia: Corbiculacea) in
north central America. American Midland Naturalist 108:74 – 80.
Hinch, S. G., Kelly, L. J., Green, R. H. 1989. Morphological variation of Elliptio complanata (Bivalvia: Unionidae) in differing
sediments of soft-water lakes exposed to acidic deposition.
Canadian Journal of Zoology 67:1895 – 1899.
Hiripi, L., Burrell, D. E., Brown, M., Assanah, P., Stanec, A., Stefano,
G. B. 1982. Analysis of monoamine accumulations in the neuronal tissues of Mytilus edulis and Anodonta cygnea (Bivalvia)III. Temperature and seasonal influences. Comparative Biochemistry and Physiology, C 71:209 – 213.
Hiscock, I. D. 1953. Osmoregulation in Australian freshwater mussels (Lamellibranchiata). I. Water and chloride exchange in
Hyridella australis (Lam.). Australian Journal of Marine and
Freshwater Research 4:317 – 329.
Hoeh, W. R., Trdan, R. J. 1984. The freshwater mussels (Pelecypoda:
Unionidae) of the upper Tittabawassee River drainage, Michigan. Malacological Review 17:97 – 98.
Hoeh W. R., Frazer, K. F., Naranjo-García, E., Trdan, R. J. 1995. A
phylogenetic perspective on the evolution of simultaneous hermaphroditism in a freshwater mussel clade (Bivalvia: Unionidae: Utterbackia). Malacological Review 28:25 – 42.
Hoke, E. 1994. A survey and analysis of the unionid mollusks of the
Elkhorn River Basin, Nebraska. Transactions of the Nebraska
Academy of Sciences 21:31 – 54.
Holopainen, I. J. 1987. Seasonal variation of survival time in anoxic
water and the glycogen content of Sphaerium corneum and Pisidium amnicum (Bivalvia: Pisidiidae). American Malacological
Bulletin 5:41 – 48.
Holopainen, I. J., Hanski, I. 1986. Life history variation in Pisidium.
Holarctic Ecology 9:85 – 98.
Holopainen, I. J., Jonasson, P. M. 1983. Long-term population dynamics and production of Pisidium (Bivalvia) in the profundal
of Lake Esrom, Denmark. Oikos 41:99 – 117.
11. Mollusca: Bivalvia
Hornbach, D. J. 1985. A review of metabolism in the Pisidiidae with
new data on its relationship with life history traits in Pisidium
casertanum. American Malacological Bulletin 3:187 – 200.
Hornbach, D. J. 1991. The influence of oxygen availability on oxygen consumption in the freshwater clam Musculium partumeium (Say) (Bivalvia: Sphaeriidae). American Malacological
Bulletin 9:39 – 42.
Hornbach, D. J., Childers, D. L. 1986. Life-history variation in a
stream population of Musculium partumeium (Bivalvia: Pisidiidae). Journal of the North American Benthological Society
5:263 – 271.
Hornbach, D. J., Childers, D. L. 1987. The effect of acidification on
life-history traits of the freshwater clam Musculium partumeium (Say, 1822) (Bivalvia: Pisidiidae). Canadian Journal of
Zoology 65:113 – 121.
Hornbach, D. J., Cox, C. 1987. Environmental influences on life history traits in Pisidium casertanum (Bivalvia: Pisidiidae): field
and laboratory experimentation. American Malacological Bulletin 5:49 – 64.
Hornbach, D. J., Way, C. M., Burky, A. J. 1980. Reproductive strategies in the freshwater sphaeriid clam, Musculium partumeium
(Say), from a permanent and a temporary pond. Oecologia
44:164 – 170.
Hornbach, D. J., Wissing, T. E., Burky, A. J. 1982. Life-history characteristics of a stream population of the freshwater clam
Sphaerium striatinum Lamarck (Bivalvia: Pisidiidae). Canadian
Journal of Zoology 60:249 – 260.
Hornbach, D. J., Wissing, T. E., Burky, A. J. 1983. Seasonal variation
in the metabolic rates and Q10 values of the fingernail clam,
Sphaerium strianum Lamark. Comparative Biochemistry and
Physiology. A76:783 – 790.
Hornbach, D. J., Way, C. M., Wissing, T. E., Burky, A. J. 1984a.
Effect of particle concentration and season on the filtration
rates of the freshwater clam, Sphaerium striatinum Lamark
(Bivalvia: Pisidiidae). Hydrobiologia 108:83 – 96.
Hornbach, D. J., Wissing, T. E., Burky, A. J. 1984b. Energy budget
for a stream population of the freshwater clam. Sphaerium striatium Lamark (Bivalvia: Pisidiidae). Canadian Journal of Zoology 62:2410 – 2417.
Hornbach, D. J., Miller, A. C., Payne, B. S. 1992. Species composition of the mussel assemblages in the upper Mississippi River.
Malacological Review 25:119 – 128.
Horne, F. R., MacIntosh, S. 1979. Factors influencing distribution of
mussels in the Blanco River of central Texas. Nautilus
94:119 – 133.
Horohov, J., Silverman, H., Lynn, J. W., Dietz, T. H. 1992. Ion transport in the freshwater zebra mussel, Dreissena polymorpha. Biological Bulletin (Woods Hole, MA) 183:297 – 303.
Houp, R. E. 1993. Observations on long-term effects of sedimentation on freshwater mussels (Mollusca: Unionidae) in the North
Fork of Red River, Kentucky. Transactions of the Kentucky
Academy of Sciences 54:93 – 97.
Hove, M. C., Neves, R. J. 1994. Life history of the endangered James
spiny mussel Pleurobema collina (Conrad, 1837) (Mollusca:
Unionidae). American Malacological Bulletin 11:29 – 40.
Hudson, R. G., Isom, B. G. 1984. Rearing of juveniles of the freshwater
mussels (Unionidae) in a laboratory setting. Nautilus 98:129– 135.
Huebner, J. D. 1982. Seasonal variation in two species of unionid
clams from Manitoba, Canada: respiration. Canadian Journal
of Zoology 60:650 – 564.
Huebner, J. D., Pynnönen, K. S. 1992. Viability of glochidia of two
species of Anodonta exposed to low pH and selected metals.
Canadian Journal of Zoology 70:2348 – 2355.
Huebner, J. D., Malley, D. F., Donkersloot, K. 1990. Population
ecology of the freshwater mussel Anodonta grandis grandis in
421
a Precambrian Shield lake. Canadian Journal of Zoology
68:1931 – 1941.
Hunter, D. R., Baily, J. F. 1992. Dreissena polymorpha (zebra mussel): colonization of soft substrata and some effects on unionid
bivalves. Nautilus 106:60 – 67.
Imlay, M. J. 1982. Use of shells of freshwater mussels in monitoring
heavy metals and environmental stresses: a review. Malacological Review 15:1 – 14.
Isom, B. G., Gooch, C. 1986. Rational and sampling designs for
freshwater mussels Unionidae in streams, large rivers, impoundments and lakes. in: Isom, B. G., Ed., Rationale for sampling
and interpretation of ecological data in the assessment of freshwater ecosystems, ASTM STP 894. American Society for Testing and Materials, Philadelphia, PA, pp. 46 – 59.
Isom, B. G., Hudson, R. G. 1982. In vitro culture of parasitic freshwater mussel glochidia. Nautilus 96:147 – 151.
Istin, M., Girard, J. P. 1970a. Dynamic state of calcium reserves in
freshwater clam mantle. Calcified Tissue Research 5:196 – 206.
Istin, M., Girard, J. P. 1970b. Carbonic anhydrase and mobilization
of calcium reserves in the mantle of lamellibranchs. Calcified
Tissue Research 5:247 – 260.
Jadhav, M. L., Lomte, V. S. 1982a. Seasonal variation in biochemical
composition of the freshwater bivalve, Lamellidens corrianus.
Rivista di Idrobiologia 21:1 – 17.
Jadhav, M. L., Lomte, V. S. 1982b. Hormonal control of carbohydrate metabolism in the freshwater bivalve, Lamellidens corrianus. Rivista di Idrobiologia. 21:27 – 36.
Jadhav, M. L., Lomte, V. S. 1983. Neuroendocrine control of midgut
gland in the bivalve, Lamellidens corrianus (Prasad) (Mollusca:
Lamellibranchiata). Journal of Advanced Zoology 4:97 – 104.
James, M. R. 1985. Distribution, biomass and production of the
freshwater mussel, Hyridella menziesi (Gray), in Lake Taupo,
New Zealand. Freshwater Biology 15:307 – 314.
Jansen, W. A., Hanson, J. M. 1991. Estimates of the number of
glochidia produced by clams (Anodonta grandis simpsoniana
Lea), attaching to yellow perch (Perca flavescens), and surviving
to various ages in Narrow Lake, Alberta. Canadian Journal of
Zoology 69:973 – 977.
Johnson, P. D., Brown, K. M. 1998. Intraspecfic life history variation
in the threatened Louisiana pearlshell mussel, Margaritifera
hembeli. Freshwater Biology 40:317 – 329.
Johnson, P. D., Brown, K. M. 2000. The importance of microhabitat
factors and habitat stability to the threatened Louisiana pearl
shell, Margaritifera hebeli (Conrad). Canadian Journal of Zoology 78:1 – 7.
Johnson, P. D., McMahon, R. F. 1998. Effects of temperature and
chronic hypoxia on survivorship of the zebra mussel (Dreissena
polymorpha) and Asian clam (Corbicula fluminea). Canadian
Journal of Fisheries and Aquatic Science 55:1564 – 1572.
Johnson, R. I. 1970. The systematics and zoogeography of the Unionidae (Mollusca: Bivalvia) of the southern Atlantic Slope Region.
Bulletin of the Museum of Comparative Zoology 140:263 – 450.
Johnson, R. I. 1972. The Unionidae (Mollusca: Bivalvia) of Peninsular Florida. Bulletin of the Florida State Museum, Biological
Sciences 16:181 – 249 addendum.
Johnson, R.I. 1998. A new mussel Potamilus metnecktayi (Bivalvia:
Unionidae) from the Rio Grande system, Mexico and Texas
with notes on Mexican Disconaias. Occasional papers on Mollusks 5(76):427 – 455.
Jokinen, E. H. 1991. The malacofauna of the acid and non-acid lakes
and rivers of the Adirondack Mountains and surrounding lowlands, New York State, USA. Verhandlungen-Internationale
Vereinigung fur Limnologie 24:2940 – 2946.
Jokela, J., Palokamgas, P. 1993. Reproductive tactics in Anodonta
clams: parental host recognition. Animal Behavior 1993:618–620.
422
Robert F. McMahon and Arthur E. Bogan
Jokela, J., Uotila, L., Taskinen, J. 1993. Effect of the castrating
trematode parasite Rhipidocotyle fennica on energy allocation
of freshwater clam Anodonta piscinalis. Functional Ecology
7:332 – 338.
Jonasson, P. M. 1984a. Decline of zoobenthos through five decades
of euthrophication in Lake Esrom. Verhandlungen Internationale Vereinigung fur Theoretische and Angewandte Limnologie 22:800 – 804.
Jonasson, P. M. 1984b. Oxygen demand and long term changes in
zoobenthos. Hydrobiologia 115:121 – 126.
Jones, H. D. 1983. Circulatory systems of gastropods and bivalves.
in: Saleuddin, A. S. M., Wilbur, K. M., Eds. The Mollusca. Vol.
5: Physiology, Part 2. Academic Press, New York, pp. 189 – 238.
Jones, H. D., Peggs, D. 1983. Hydrostatic and osmotic pressure in the
heart and pericardium of Mya arenaria and Anodonta cygnea.
Comparative Biochemistry and Physiology 76A:381 – 385.
Jones, R. G., Davis, W. L. 1982. Calcium containing lysosomes in the
outer mantle epithelial cells of Amblema, a freshwater mollusc.
Anatomic Record 203:337 – 343.
Joosse, J., Geraerts, W. P. M. 1983. Endocrinology. in: Saleuddin, A.
S. M., Wilbur, K. M., Eds. The Mollusca, Vol. 4: Physiology,
Part 1. Academic Press, New York, pp. 318 – 406.
Jørgensen, C. B. 1996. Bivalve filter feeding revisited. Marine Ecology Progress Series 142:287 – 302.
Jørgensen, C. B., Kiorboe, T., Mohlenberg, F., Riisgard, H. U. 1984.
Ciliary and mucus-net filter feeding, with special reference to
fluid mechanical characteristics. Marine Ecology Progress Series
15:283 – 292.
Kasprzak, K. 1986. Role of the Unionidae and Sphaeriidae (Mollusca, Bivalvia) in the eutrophic Lake Zbechy and its outflow.
Internationale
Revue
der
Gesamten
Hydrobiologie
71:315 – 334.
Kat, P. W. 1982. Shell dissolution as a significant cause of mortality
for Corbicula fluminea (Bivalvia: Corbiculidae) inhabiting
acidic waters. Malacological Review 15:129 – 134.
Kat, P. W. 1984. Parasitism and the Unionacea (Bivalvia). Biological
Review 59:189 – 207.
Kat, P. W. 1985. Convergence in bivalve conchiolin layer microstructure. Malacological Review 18:97 – 106.
Kat, P. W., Davis, G. M. 1984. Molecular genetics of peripheral population of Nova Scotian Unionidae (Mollusca: Bivalvia). Biological Journal of the Linnean Society 22:157 – 185.
Kays, W. T., Silverman, H., Dietz, T. H. 1990. Water channels and
water canals in the gill of the freshwater mussel, Ligumia subrostrata: ultrastructure and histochermistry. Journal of Experimental Zoology 254:256 – 269.
Keen, M., Casey, R. 1969. Family Corbiculidae Gray, 1847. in:
Moore, R. C., Ed. Treatise on invertebrate paleontology. Part
N: Mollusca. The Geological Society of America., Boulder, CO,
pp. 664 – 669.
Keen, M., Dance, P. 1969. Family Pisidiidae, Gray, 1857. in: Moore,
R. C., Ed. Treatise on invertebrate paleontology. Part N: Mollusca. The Geological Society of America, Boulder, CO, pp.
669 – 670.
Kennedy, V. S., Blundon, J. A. 1983. Shell strength in Corbicula sp.
(Bivalvia: Corbiculacea) from the Potomac River, Maryland.
Veliger 26:22 – 25.
Khan, H. R., Aston, M. L., Saleuddin, A. S. M. 1986. Fine structure
of the kidneys of osmotically stressed Mytilus, Mercenaria, and
Anodonta. Canadian Journal of Zoology 64:2779 – 2787.
Kilgour, B. W., Mackie, G. L. 1988. Factors affecting the distribution
of sphaeriid bivalves in Britannia Bay of the Ottawa River. Nautilus 102:73 – 77.
King, C. A., Langdon, C. J., Counts, C. L., III. 1986. Spawning and
early development of Corbicula fluminea (Bivalvia: Corbiculi-
dae) in laboratory culture. American Malacological Bulletin
4:81 – 88.
Kirk, S. G., Layzer, J. B. 1997. Induced metamorphosis of freshwater
mussel glochidia on nonfish host. Nautilus 110:102 – 106.
Kiss, A., Pekli, J. 1988. On the growth of Anodonta woodiana (Lea
1834) (Bivalvia: Unionacea). Bulletin of the University of Agricultural Sciences, Gödöllö 1:119 – 124.
Kovolack, W. P., Dennis, S. D., Bates, J. M. 1986. Sampling effort required to find rare species of freshwater mussels. in: Isom, B.
G., Ed. Rationale for sampling and interpretation of ecological
data in the assessment of freshwater ecosystems, ASTM STP
894. American Society for Testing and Materials, Philadelphia,
PA, pp. 34 – 45.
Kraemer, L. R. 1978. Discovery of two distinct kinds of statocysts in
freshwater bivalved mollusks: some behavioral implications.
Bulletin of the American Malacological Union pp. 24 – 28.
Kraemer, L. R. 1981. The osphradial complex of two freshwater bivalves: Histological evaluation and functional context. Malacologia 20:205 – 216.
Kraemer, L. R., Galloway, M. L. 1986. Larval development of
Corbicula fluminea (Müller) (Bivalvia: Corbiculacea): an appraisal of its heterochrony. American Malacological Bulletin
4:61 – 79.
Kraemer, L. R., Swanson, C., Galloway, M., Kraemer, R. 1986. Biological basis of behavior in Corbicula fluminea, II. Functional
morphology of reproduction and development and review of evidence for self-fertilization. American Malacological Bulletin,
Special Edition. No. 2:193 – 201.
Kryger, J., Riisgård, H. U. 1988. Filtration rate capacities in 6 species
of European freshwater bivalves. Oecologia 77:34 – 38.
LaRocque, A. L. 1967a. Pleistocene Mollusca of Ohio. Part 2. State
of Ohio Division of Geological Survey, Department of Natural
Resources, Columbus. pp. 113 – 356.
LaRocque, A. L. 1967b. Pleistocene Mollusca of Ohio. Part1. State
of Ohio Department of Natural Resources, Division of Geological Survey, Columbus. pp. 1 – 111.
Lauritsen, D. D. 1986a. Assimilation of radiolabeled algae by Corbicula. American Malacological Bulletin, Special Edition No.
2:219 – 222.
Lauritsen, D. D. 1986b. Filter-feeling in Corbicula fluminea and its
effect on seston removal. Journal of the North American Bentholoigical Society 5:165 – 172.
Laycock, G. 1983. Vanishing naiads. Audubon 85:26 – 28.
Leff, D. S., Burch, J. L., McArthur, J. V. 1990. Spatial distribution,
seston removal, and potential competitive interactions of the bivalves Corbicula fluminea and Elliptio complanata in a coastal
plain stream. Freshwater Biology 24:409 – 416.
Lei, J., Payne, B. S., Wang, S. Y. 1996. Filtration dynamics of the zebra mussel, Dreissena polymorpha. Canadian Journal of Fisheries and Aquatic Science 53:29 – 37.
Lewis, J. B. 1984. Comparative respiration in two species of freshwater
unionid mussels (Bivalvia). Malacological Review 17:101 – 102.
Lewis, J. B., Riebel, P. N. 1984. The effect of substrate on burrowing
in freshwater mussels (Unionidae). Canadian Journal of Zoology 62:2023 – 2025.
Lomte, V. S., Jadhav, M. L. 1981a. Effect of desiccation on the neurosecretory activity of the freshwater bivalve, Parreysia corrugata. Science and Culture 47:437 – 438.
Lomte, V. S., Jadhav, M. L. 1981b. Neuroendocrine control of osmoregulation in the freshwater bivalve, Lamellidens corrianus.
Journal of Advanced Zoology 2:102 – 108.
Lomte, V. S., Jadhav, M. L. 1982a. Effects of toxic compounds on
oxygen consumption in the freshwater bivalve, Corbicula regularis (Prime, 1860). Comparative Physiology and Ecology
7:31 – 33.
11. Mollusca: Bivalvia
Lomte, V. S., Jadhav, M. L. 1982b. Respiratory metabolism in the
freshwater mussel Lamellidens corrianus. Marathwada University Journal of Science 21:87 – 90.
Lomte, V. S., Jadhav, M. L. 1982c. Biochemical composition of the
freshwater bivalve, Lamellidens corrianus (Prasad, 1922).
Rivista di Idrobiologia 21:19 – 25.
Lopez, G. R., Holopainen, I. J. 1987. Interstitial suspension-feeding
by Pisidium sp. (Pisididae: Bivalvia): a new guild in the lentic
benthos? American Malacological Bulletin 5:21 – 30.
Lynn, J. W. 1994. The ultrastructure of the sperm and motile spermatozeugmata released from the freshwater mussel Anodonta
grandis (Mollusca: Bivalvia, Unionoidae). Canadian Journal of
Zoology 72:1452 – 1461.
Lydeard, C., Mulvey, M., Davis, G. M. 1996. Molecular systematics
and evolution of reproductive traits of North American freshwater unionacean mussels (Mollusca: Bivalvia) as inferred from
16S rRNA gene sequences. Philosophical Transactions of the
Royal Society of London B, 351:1593 – 1603.
Machodo, J. F., Ferreira, K. G., Ferreira, H. G., Fernandes, P. L. 1990.
The acid – base balance of the outer mantle epithelium of Anodonta cygnea. Journal of Experimental Biology 150:159 – 169.
Machado, J., Marvo, R., Ferreira, C., Moura, G., Ries, M., Coimbra,
C. 1994. Study on mucopolysaccharides as a shell component
of Anodonta cygnea. Bulletin de l’Institut Océanographique,
Monaco 14:141 – 150.
Machena, C., Kautsky, N. 1988. A quantitative diving survey of benthic vegetation and fauna in Lake Kariba, a tropical man-made
lake. Freshwater Biology 19:1 – 14.
MacIsaac, H. J. 1996. Potential abiotic and biotic impacts of zebra
mussels on the inland waters of North America. American Zoologist 36:287 – 299.
MacIsaac, H. J., Johannsson, O. E., Ye, J., Sprules, W. G., Leach, J.
H., McCorquodale, J. A., Grigorovich, I. A. 1999. Filtering impacts of an introduced bivalve (Dreissena polymorpha) in a
shallow lake: application of a hydrodynamic model. Ecosystems
2:338 – 350.
Mackie, G. L. 1978. Are sphaeriid clams ovoviparous or viviparous?
Nautilus 92:145 – 147.
Mackie, G.L. 1979. Growth dynamics in natural populations of
Sphaeriidae clams (Sphaerium, Musculium, Pisidium). Canadian
Journal of Zoology 57:441 – 456.
Mackie, G. L. 1984. Bivalves. in: Tompa, A. S., Verdonk, N. H., van
der Biggelaar, J. A. M. Eds. The Mollusca. Vol. 7: Reproduction. Academic Press, New York, pp. 351 – 418.
Mackie, G. L., Flippance, L. A. 1983a. Life history variations in two
populations of Sphaerium rhombiodeum (Bivalvia: Pisidiidae).
Canadian Journal of Zoology 61:860 – 867.
Mackie, G. L., Flippance, L. A. 1983b. Intra and interspecific variation in calcium content of freshwater Mollusca in relation to
calcium content of the water. Journal of Molluscan Studies
49:204 – 212.
Mackie, G. L., Schloesser, D. W. 1996. Comparative biology of zebra
mussels in Europe and North America: an overview. American
Zoologist 36:244 – 258.
Mackie, G. L., Topping, J. M. 1988. Historical changes in the
unionid fauna of the Sydenham River Watershed and downstream changes in shell morphometrics of three common
species. Canadian Field Naturalist 102:617 – 626.
Mackie, G. L., Quadri, S. U., Reed, R. M. 1978. Significance of litter
size in Musculium securis (Bivalvia: Sphaeriidae). Ecology
59:1069 – 1074.
Mackie, G. L., White, D. S., Zdeba, T. W. 1980. A guide to the freshwater mollusks of the Laurentian Great Lakes with special emphasis on the genus, Pisidium. EPA-600 / 3-80-068. United
States Environmental Protection Agency, Duluth, MN.
423
Mackie, G. L., Gibbons, W. N., Muncaster, B. W., Gray, I. M. 1989.
The zebra mussel, Dreissena polymorpha: a synthesis of European experience and a preview for North America. 0-77295647-2. Water Resources Branch, Ontario Ministry of the Environment, Ont., Canada.
Mäkelä, T. P., Oikari, A. O. J. 1992. The effects of low water pH on
the ionic balance in freshwater mussel Anodonta anatina L. Annales Zoologici Fennici 29:169 – 175.
Malley, D. F., Huebner, J. D., Donkersloot, K. 1988. Effects on ionic
composition of blood and tissues of Anodonta grandis grandis
(Bivalvia) of an addition of aluminum and acid to a lake.
Archives of Environmental Contamination and Toxicology
17:479 – 491.
Malloy, D. P., Karatayev, A. Y., Burlakova, L. E., Kurandina, D. P.,
Laruelle, F. 1997. Natural enemies of zebra mussels, and ecological competitors. Reviews in Fisheries Science 5:27 – 97.
Mani, V. U., Rao, P. V., Reddy, K. S., Indira, K., Rajendra, W. 1993.
Bioaccumulation and clearance of ambient ammonia by the
freshwater water mussel, Lamellidens marginalis (L.). Indian
Journal of Comparative Animal Physiology 11:83 – 88.
Marsden, J. E., Spidle, A. P., May, B. 1996. Review of genetic studies
of Dreissena sp. American Zoologist 36:259 – 270.
Marsh, P. C. 1985. Secondary production of introduced Asiatic clam,
Corbicula fluminea, in a central Arizona canal. Hydrobiologia
124:103 – 110.
Martin, W. A. 1983. Execretion. in: Saleuddin, A. S. M., Wilbur, K.
M. Eds. The Mollusca. Vol. 5: Physiology, Part 2. Academic
Press. New York, pp. 353 – 405.
Massabuau, J-C., Burtin, B., Wheatly, M. 1991. How is O2 consumption maintained independent of ambient oxygen in mussel, Anodonta cygnea? Respiration Physiology 83:103 – 114.
Mathiak, H. A. 1979. A river survey of the unionid mussels of Wisconsin 1973 – 1977. Sand Shell Press. Horicon, WIS.
Matisoff, G., Fisher, J. B., Matis, S. 1985. Effect of microinvertebrates on the exchange of solutes between sediments and freshwater. Hydrobiologia 122:19 – 33.
Matsushima, O., Kado, Y. 1982. Hyperosmoticity of the mantle fluid
in the freshwater bivalve, Anodonta woodiana. Journal of Experimental Biology 221:379-381.
Matsushima, O., Sakka, F. and Kado, Y. 1982. Free amino acid involved in intracellular osmoregulaion in the clam, Corbicula.
Journal of Science of Hiroshima University, Series B, Division 1
30:213 – 219.
Mattice, L. S. 1979. Interactions of Corbicula sp. with power plants.
in; J. C. Britton, Ed. Proceedings, first international Corbicula
symposium. Texas Christian University Research Foundation,
Fort Worth, TX, pp. 119 – 138.
Mattice, J. S., McLean, R. B., Burch, M. B. 1982. Evaluation of
short-term exposure to heated water and chlorine for control of
the Asiatic clam (Corbicula fluminea). Oak Ridge National Laboratory, Environmental Science Division, Publication No. 1748.
U.S. National Technical Information Service, Department of
Commerce, Springfield, VA, 33 pp.
Matthews, M. A., McMahon, R. F. 1999. Effects of temperature
and temperature acclimation on survival of zebra mussels
(Dreissena polymorpha) and Asian clams (Corbicula fluminea)
under extreme hypoxia. Journal of Molluscan Studies 65:
317 – 325.
Mazak, E. J., MacIsaac, H. J., Servos, M. R., Hesslein, R. 1997. Influence of feeding habits on organochlorine contaminant accumulation in waterfowl of the Great Lakes. Ecological Applications 7:1133 – 1143.
McCorkle, S., Shirley, T. C., Dietz, T. H. 1979. Rhythms of activity
and oxygen consumption in the common pond clam, Ligumia
subrostrata (Say). Canadian Journal of Zoology 57: 1960 – 1964.
424
Robert F. McMahon and Arthur E. Bogan
McCorkle-Shiley, S. 1982. Effects of photoperiod on sodium flux in
Corbicula fluminea (Mollusca: Bivalvia). Comparative Biochemistry and Physiology. A71:325 – 327.
McKee, P. M., Mackie, G. L. 1980. Dessication resistance in
Sphaerium occidentale and Musculium securis (Bivalvia:
Sphaeriidae) from a temporary pond. Canadian Journal of
Zoology 58:1693 – 1696.
McKillop, W. B., Harrison, A. D. 1982. Hydrobiological studies of
the eastern Lesser Antillean Islands. VII. St. Lucia: behavioural
drift and other movements of freshwater marsh molluscs.
Archiv für Hydrobiologie 94:53 – 69.
McLeod, M. J. 1986. Electrophoretic variation in North America
Corbicula. American Malacological Bulletin, Special ed. No.
2:125 – 132.
McMahon, R. F. 1979a. Response to temperature and hypoxia in the
oxygen consumption of the introduced Asiatic freshwater clam
Corbicula fluminea (Müller). Comparative Biochemistry and
Physiology A63:383 – 388.
McMahon, R. F. 1979b. Tolerance of aerial exposure in the Asiatic
freshwater clam, Corbicula fluminea (Müller). in: Britton, J. C.
Ed. Proceedings, first international Corbicula symposium. Texas
Christian University Research Foundation, Fort Worth, TX,
pp. 227 – 241.
McMahon, R. F. 1983a. Ecology of an invasive pest bivalve, Corbicula. in: Russell-Hunter, W. D. Editor. The Mollusca. Vol. 6:
Ecology. Academic Press, New York, pp. 505 – 561.
McMahon, R. F. 1983b. Physiological ecology of freshwater pulmonates. in: Russell-Hunter, W. D. Ed. The Mollusca. Vol. 6:
Ecology. Academic Press, New York, pp. 360 – 430.
McMahon, R. F. 1996. The physiological ecology of the zebra mussel, Dreissena polymorpha, in North America. American Zoologist 36:339 – 363.
McMahon, R. F. 1999. Invasive characteristics of the freshwater bivalve, Corbicula fluminea. in: Claudi, R., Leach, J. H. Eds.
Nonindigenous freshwater organisms: vectors, biology and impacts. Lewis Publishers, Boca Raton, FL, pp. 315 – 343.
McMahon, R. F., Lutey, R. W. 1988. Field and laboratory studies of
the efficacy of poly[oxyethylene(dimethyliminio)ethylene(dimenthyliminio)ethylene dichloride] as a biocide against the
Asian clam, Corbicula fluminea. in: Proceedings: service water
reliability improvement seminar. Electric Power Research Institute, Palo Alto, CA, pp. 61 – 72.
McMahon, R. F., Williams, C. J. 1984. A unique respiratory response to emersion in the introduced Asian freshwater clam
Corbicula fluminea (Müller) (Lamellibranchia: Corbiculacea).
Physiological Zoology 57:274 – 279.
McMahon, R. F., Williams, C. J. 1986a. A reassessment of growth
rate, life span, life cycles and population dynamics in a natural
population and field caged individuals of Corbicula fluminea
(Müller). American Malacological Bulletin, Special ed. No.
2:151 – 166.
McMahon, R. F., Williams, C. J. 1986b. Growth, life cycle, upper
thermal limit and downstream colonization rates in a natural
population of the freshwater bivalve mollusc, Corbicula fluminea (Müller) receiving thermal effluents. American Malacological Bulletin, Special ed. No. 2:231 – 239.
McMahon, R. F., Ussary, T. A., Clarke, M. 1993. Use of emersion as
a zebra mussel control method, Technical Report EL-93-1, U.S.
Army Corps of Engineers, Washington, DC.
Medler, S., Silverman, H. 1997. Functional organization of intrinsic
gill muscles in zebra mussels, Dreissena polymorpha (Mollusca:
Bivalvia), and response to transmitters in vivo. Invertebrate Biology 116:200 – 212.
Michaelidis, B., Anthanasiadou, P. 1994. Effect of reduced oxygen
tension on the heart rate and the kinetic properties of glycolytic
key enzymes PFK, PK and glycogen phosphorylase from the
freshwater mussel, Anodonta cygnea. Comparative Biochemistry and Physiology B108:165 – 172.
Miller, A. C., Payne, B. S. 1988. The need for quantitative sampling
to characterize demography and density of freshwater mussel
communities. American Malacological Bulletin 6:49 – 54.
Miller, A. C., Rhodes, L., Tipit, R. 1984. Changes in the naiad fauna
of the Cumberland River below Lake Cumberland in central
Kentucky. Nautilus 98:107 – 110.
Miller, A. C., Payne, B. S., Aldridge, D. W. 1986. Characterization of
a bivalve community in the Tangipahoa River, Mississippi. Nautilus 100:18 – 23.
Miller, A. C., Whiting, R., Wilcox, D. B. 1989. An evaluation of a
skimmer dredge for collecting freshwater mussels. Journal of
Freshwater Ecology 5:151 – 154.
Miller, A. C., Payne, B. S., Tippet, R. 1992. Characterization of a
freshwater mussel (Unionidae) community immediately downriver of Kentucky Lock and Dam in the Tennessee River. Transactions of the Kentucky Academy of Sciences 53:154 – 161.
Miller, A. C., Lei, J., Tom J., 1994. Shell strength of the non-indigenous zebra mussel Dreissena polymorpha (Pallas) in comparison
to two other freshwater bivalve species. Veliger 37:319 – 321.
Mills, E. L., Rosenberg, G., Spidle, A. P., Ludyanskiy, M., Pligin, Y.,
May, B. 1996. A review of the biology and ecology of the
quagga mussel (Dreissena bugensis), a second species of freshwater dreissenid introduced to North America. American Zoologist 36:271 – 286.
Mills, H. B., Starrett, W. C., Belrose, F. C. 1966. Man’s effect on the
fish and wildlife of the Illinois River. Biological Notes (Illinois
Natural History Survey) No. 57:24 pp.
Mitchell, R. D. 1955. Anatomy, life history, and evolution of the
mites parasitizing freshwater mussels. Miscellaneous Publications Museum of Zoology University of Michigan 89:1 – 28.
Morton, B. 1982. Some aspects of the population structure and sexual strategy of Corbicula cf. fluminails (Bivalvia: Corbiculacea)
from the Pearl River. Peoples Republic of China. Journal of
Mollusca Studies 48:1 – 23.
Morton, B. 1983. Feeding and digestion in Bivalvia. in: Saleuddin, A.
S. M., Wilbur, K. M. Eds. The Mollusca. Vol. 5: Physiology,
Part 2. Academic Press, New York, pp. 65 – 147.
Morton, B. 1985. The population dynamics, reproductive strategy
and life history tactics of Musculium lacustre (Bivalvia: Pisidiidae) in Hong Kong. Journal of Zoology (London) (A)
207:581 – 603.
Morton, B. 1986. The population dynamics and life history tactics of
Pisiduim clarkeanum and P. annandalei (Bivalvia: Pisidiidae)
sympatric in Hong Kong. Journal of Zoology (London) (A)
210:427 – 449.
Morton, B., Tong, K. Y. 1985. The salinity tolerance of Corbicula
fluminea (Bivalvia: Corbiculoidea) from Hong Kong. Malacological Review 18:91 – 95.
Nagabhushanam, R., Lomte, V. S. 1981. Observations on the relationship between neurosecretion and periodic activity in the
mussel, Parreysia corrugata. Indian Journal of Fisheries
28:261 – 265.
Nalepa, T. F., Gauvin, J. M. 1988. Distribution, abundance, and biomass of freshwater mussels (Bivalvia: Unionidae) in Lake St.
Clair. Journal of Great Lakes Research 14:411 – 419.
Nalepa, T. F., Gardner, W. S., Malczyk, J. M. 1991. Phosphorus cycling by mussels (Unionidae: Bivalvia) in Lake St. Clair. Hydrobiologia 219:239 – 250.
Narain, A. S., Singh, K. 1990. Unionid rectal structure in relation to
function and with particular reference to the piercing of heart
by gut. Proceedings of the National Academy of Science, India
60:245 – 252.
11. Mollusca: Bivalvia
Negus, C . 1966. A quantitative study of growth and production of
unionid mussels in the River Thames at Reading. Journal of Animal Ecology 35:513 – 532.
Nelson, D. J., Scott, D. C. 1962. Role of detritus in the productivity
of a rock – outcrop community in a Piedmont stream. Limnology and Oceanography 7:396 – 413.
Neves, R. J. 1993. A state-of-the-unionids address. in: Cummings, K.
S., Buchanan, A. C., Koch, L. M. Eds. Conservation and management of freshwater mussels. Proceedings of a Symposium,
1992, Upper Mississippi River Conservation Committee, Rock
Island, IL, pp. 2 – 9.
Neves, R. J. 1997. A national strategy for the conservation of native
freshwater mussels. in: Cummings, K. S., Buchanan, A. C.,
Mayer, C. A., Naimo, T. J. Eds. Conservation and management
of freshwater mussels II: initiatives for the future. Proceedings
of a Symposium, 1995, Upper Mississippi River Conservation
Committee, Rock Island, IL, pp. 1 – 10.
Neves, R. J., Moyer, S. N. 1988. Evaluation of techniques for age determination of freshwater mussels (Unionidae). American Malacological Bulletin 6:179 – 188.
Neves, R. J., Zale, A. V. 1982. Freshwater mussels (Unionidae) of Big
Moccasin Creek, Southwestern Virginia. Nautilus 96:52 – 54.
Neves, R. J., Weater, L. R., Zale, A. V. 1985. An evaluation of host
fish suitability for glochidia of Villosa vanuxemi and V. nebulosa (Pelecypoda: Unionidae). American Midland Naturalist
113:13 – 18.
Neves, R. J., Bogan, A. E., Williams, J. D., Ahlstedt, A. S., Hartfield,
P. W. 1997. Status of aquatic mollusks in the southeastern
United States: a downward spiral of diversity. in: Benz, G. W.,
Collins, D. E. Eds. Aquatic fauna in peril: the southeastern perspective, Special Publication 1, Southeast Aquatic Research Institute, Chattanooga, TN, pp. 43 – 85.
New York Sea Grant. 1999. North American range of the zebra mussel. Dreissena! 10:8 – 9.
Nichols, S. J. 1996. Variations in the reproductive cycle of Dreissena
polymorpha in Europe, Russia, and North America. American
Zoologist 36:311 – 325.
Nichols, S. J. and Black, M.G. 1994. Identification of larvae: the zebra mussel (Dreissena polymorpha), quagga mussel (Dreissena
rosteriformis bugensis), and Asian clam (Corbicula fluminea).
Canadian Journal of Zoology 72:406 – 417.
Oesch, R. D. 1984. Missouri naiades. A guide to the mussels of
Missouri. Missouri Department of Conservation, Jefferson
City, MO.
Okland, K. A., Kuiper, J. G. J. 1982. Distribution of small mussels
(Sphariidae) in Norway, with notes on their ecology. Malacologia 22:469 – 477.
Ortmann, A. E. 1909. The destruction of the freshwater fauna in
western Pennsylvania. Proceedings of the American Philosophical Society 48:90 – 110.
Ortmann, A. E. 1918. The nayades (freshwater mussels) of the Upper
Tennessee drainage. With notes on synonymy and distribution.
Proceedings of the American Philosophical Society 57:521 – 626.
Paloumphis, A. A., Starrett, W. C. 1960. An ecological study of benthic organisms in three Illinois River flood plain lakes. American Midland Naturalist 64:406 – 435.
Panha, S. 1993. Glochidiosis and juvenile production in a freshwater
pearl mussel, Chamberlainia hainesiana. Invertebrate Reproduction and Development 24:157 – 160.
Park, J., Okino, T. 1994. Seasonal distribution and growth of
Sphaerium (Musculium) japonicum (Bivalvia; Sphaeriidae) in
Lake Suwa, Japan. Journal of the Faculty of Science, Shinsiu
University 29:49 – 56.
Parker, R. S., Hackney, C. T., Vidrine, M. F. 1984. Ecology and reproductive strategy of a south Louisiana freshwater mussel,
425
Glebula rotundata (Lamarck) (Unionidae: Lampsilini). Freshwater Invertebrate Biology 3:53 – 58.
Parmalee, P. W. 1967. The freshwater mussels of Illinois. Illinois
State Museum Popular Science Series 8:1 – 108.
Parmalee, P. W. 1988. A comparative study of late prehistoric and
modern molluscan faunas of the Little Pigeon River System,
Tennessee. American Malacological Bulletin 6:165 – 178.
Parmalee, P. W., Bogan, A. E. 1998. The freshwater mussels of Tennessee. The University of Tennessee Press, Knoxville, TN.
Parmalee, P. W., Hughes, M. H. 1993. Freshwater mussels (Mollusca:
Pelecypoda: Unionidae) of Tellico Lake: Twelve years after impoundment of the Little Tennessee River. Annals of the Carnegie
Museum 62:81 – 93.
Parmalee, P. W., Klippel, W. E. 1982. A relic population of Obovaria
retusa in the middle Cumberland River, Tennessee. Nautilus
96:30 – 32.
Parmalee, P. W., Klippel, W. E. 1984. The naiad fauna of the Tellico
River, Monroe County, Tennessee. American Malacological Bulletin 3:41 – 44.
Parmalee, P. W., Klippel, W. E., Bogan, A. E. 1982. Aboriginal and
modern freshwater mussel assemblages (Pelecypoda: Unionidae)
from the Chickamauga Reservoir, Tennessee. Brimleyana 8:
75 – 90.
Paterson, C. G. 1983. Effect of aggregation on the respiration rate of
the freshwater unionid bivalve, Elliptio complanata (Solander).
Freshwater Invertebrate Biology 2:139 – 146.
Paterson, C. G. 1984. A technique for determining apparent selective
filtration in the fresh-water bivalve Elliptio companata (Lightfoot). Veliger 27:238 – 241.
Paterson, C. G. 1985. Biomass and production of the unionid,
Elliptio companata (Lightfoot) in an old reservoir in New
Brunswick, Canada. Freshwater Invertebrate Biology 4:
201 – 207.
Paterson C. G., Cameron, I. F. 1985. Comparative energetics of two
populations of the unionid, Anodonta cataracta (Say). Freshwater Invertebrate Biology 4:79 – 90.
Paulkstis, G. L., Janzen, F. J., Tucker, J. K. 1996. Response of aerially-exposed zebra mussels (Dreissena polymorpha) to subfreezing temperatures. Journal of Freshwater Ecology 11:513 – 519.
Payne, B. S., Miller, A. C. 1987. Effects of current velocity in the
freshwater bivalve Fusconaia ebena. American Malacological
Bulletin 5:177 – 179.
Payne, B. S., Miller, A. C. 1989. Growth and survival of recent recruits
to a population of Fusconaia ebena (Bivalvia: Unionidae) in the
lower Ohio River. American Midland Naturalist 121:99 – 104.
Prezant, R. S., Charlermwat, K. 1984. Flotation of the bivalve Corbicula fluminea as means of dispersal. Science 225:1491 – 1493.
Price, R. E., Knight, L. A., Jr. 1978. Mercury, cadmium, lead and arsenic in sediments, plankton, and clams from Lake Washington
and Sardis Reservoir, Mississippi, October 1975 – May 1976.
Pesticides Monitoring Journal 11:182 – 189.
Prokopovich, N. P. 1969. Deposition of clastic sediments by clams.
Sedimentary Petrology 39:891 – 901.
Pugsley, C. W., Hebert, P. D. N., Wood, G. W., Brotea, G., Obal,
T. W. 1985. Distribution of contaminants in clams and sediments from the Huron – Erie corridor. I-PCBs and octachlorostyrene. Journal of Great Lakes Research 11:275 – 289.
Pynnönen, K. 1991. Influence of aluminum and H on the electrolyte
homeostatsis in the Unionidae, Anodonta anatina L. and Unio
pictorum L. Archives of Environmental Contamination and
Toxicology 20:218 – 255.
Quigley, M. A., Gardiner, W. S., Gordon, W. M. 1992. Metabolism
in the zebra mussel (Dreissena polymorpha) in Lake St. Clair of
the Great Lakes. in: Nalepa, T. F., Schloesser, D. W. Eds. Lewis
Publishers, Boca Raton, FL, pp. 295 – 306.
426
Robert F. McMahon and Arthur E. Bogan
Ram, J. L., McMahon, R. F. 1996. Introduction: the biology, ecology,
and physiology of zebra mussels. American Zoologist
36:239 – 243.
Ram, J. L., Fong, P. F., Garton, D. W. 1996. Physiological aspects of
zebra mussel reproduction: maturation, spawning and fertilization. American Zoologist 36:326 – 338.
Rao, K. R., Muley, S. D., Mane, U. H. 1987. Impact of some ecological factors on cholesterol content from soft tissues of freshwater
bivalve Indonaia caeruleus. Journal of Hydrobiology 3:67 – 69.
Reid, R. G. B., McMahon, R. F., Foighil, D. O., Finnigan, R. 1992.
Anterior inhalant currents and pedal feeding in bivalves. Veliger
35:93 – 104.
Rhoads, S. N. 1899. On a recent collection of Pennsylvanian mollusks from the Ohio River system below Pittsburgh. Nautilus
12:133 – 138.
Ricciardi, A., Serrouya, R., Whoriskey, F. G. 1995. Aerial exposure
tolerance of zebra and quagga mussels (Bivalvia: Dreissenidae):
implications for overland dispersal. Canadian Journal of Fisheries and Aquatic Science 52:470 – 477.
Ricciardi, A., Whoriskey, F. G., Rasmussen, J. B. 1997. The role of
zebra mussel (Dreissena polymorpha) in structuring macroinvertebrate communities on hard substrata. Canadian Journal of
Fisheries and Aquatic Science 54:2596 – 2608.
Robinson, J. V., Wellborn, G. A. 1988. Ecological resistance to the
invasion of a freshwater clam, Corbicula fluminea: fish predation effects. Oecologia 77:445 – 452.
Rocha, E., Azevedo, C. 1990. Ultrastructural study of the spermatogenesis of Anodonta cygnea L. (Bivalvia, Unionidae). Invertebrate Reproduction and Development 18:169 – 176.
Rooke, J. B., Mackie, G. L. 1984a. Mollusca of six low-alkalinity
lakes in Ontario. Canadian Journal of Fisheries and Aquatic
Science 41:777 – 782.
Rooke, J. B., Mackie, G. L. 1984b. Growth and production of three
species of molluscs in six low alkalinity lakes in Ontario,
Canada. Canadian Journal of Zoology 62:1474 – 4178.
Rooke, J. B., Mackie, G. L. 1984c. Laboratory studies of the effects
of Mollusca on alkalinity of their freshwater environment.
Canadian Journal of Zoology 62:793 – 797.
Rosenthal, R. J. 1969. A method of tagging mollusks underwater.
Veliger 11:288 – 289.
Ruppert, E. E., Barnes, R. D. 1994. Invertebrate zoology, 6th ed.,
Saunders College Publishing, Philadelphia, PA.
Russell-Hunter, W. D., Buckley, D. F. 1983. Actuarial bioenergetics of
nonmarine molluscan productivity. in: Russell-Hunter, W. D.
Ed. The Mollusca. Vol. 6: Ecology. Academic Press, New York,
pp. 464 – 503.
Sahib, I. K. A., Narasimhamurthy, B., Sailatha, D., Begum, M. R.,
Prasad, K. S., Roa, K. V. R. 1983. Orientation in the glycolytic
potentials of carbohydrate metabolism in the selected tissues of
the aestivating freshwater pelecypod, Lamellidens marginalis
(Lamarck). Comparative Physiology and Ecology 8:180 – 184.
Saintsing, D. G., Dietz, T. H. 1983. Modification of sodium transport
in freshwater in mussels by prostaglandins, cyclic AMP and 5hydroxytryptamine: effects of inhibitors of prostaglandin synthesis. Comparative Biochemistry and Physiology C76: 285 – 290.
Saleuddin, A. S. M., Petit, H. P. 1983. The mode of formation and
the structure of the periostracum. in: Saleuddin, A. S. M.,
Wilbur, K. M. Eds. The Mollusca. Vol. 4: Physiology, Part 1:
Academic Press, New York, pp. 199 – 234.
Salmon, A., Green, R. H. 1983. Environment determinants of
unionid clam distribution in the Middle Thames River, Ontario.
Canadian Journal of Zoology 61:832 – 838.
Schloesser, D. W., Nalepa, T. F., Mackie, G. L. 1996. Zebra mussel
infestation of unionid bivalves (Unionidae) in North America.
American Zoologist 36:300 – 310.
Schloesser, D. W., Smithee, R. D., Longton, G. D., Kovalak, W. P.
1997. Zebra mussel induced mortality of unionids in firm substrata of western Lake Erie and a habitat for survival. American
Malacological Bulletin 14:67 – 74.
Servos, M. R., Rooke, J. B., Mackie, G. L. 1985. Reproduction of selected Mollusca in some low alkalinity lakes in south-central
Ontario. Canadian Journal of Zoology 63:511 – 515.
Sheldon, F., Walker, K. F. 1989. Effects of hypoxia on oxygen consumption by two species of freshwater mussel (Unionacea;
Hyriidae) from the River Murray. Australian Journal of Marine
and Freshwater Research 40:491 – 499.
Sibly, R. M., Calow, P. 1986. Physiological ecology of animals: an
evolutionary approach. Blackwell, London.
Sickel, J. B. 1986. Corbicula population mortalities: factors influencing population control. American Malacological Bulletin, Special Edition. No. 2:89 – 94.
Silverman, H., Steffens, W. L., Dietz, T. H. 1983. Calcium concretions in the gills of a freshwater mussel serve as a calcium reservoir during periods of hypoxia. Journal of Experimental Zoology 227:177 – 189.
Silverman, H., Steffens, W. L., Dietz, T. H. 1985. Calcium from extracellular concretions in the gills of freshwater unionid mussels
is mobilized during reproduction. Journal of Experimental Zoology 236:137 – 147.
Silverman, H., Kays, W. T., Dietz, T. H. 1987. Maternal calcium contribution of glochidial shells in freshwater mussels (Eulamellibranchia: Unionidae). Journal of Experimental Biology 242:
137 – 146.
Silverman, H., Silby, L. D., Steffens, W. L. 1988. Calmodulin-like calcium-binding protein identified in the calcium-rich mineral deposits from freshwater mussel gills. Journal of Experimental
Zoology 247:224 – 231.
Silverman, H., Richard, P. E., Goddard, R. H., Dietz, T. H. 1989. Intracellular formation of calcium concretions by phagocytic cells
in freshwater mussels. Canadian Journal of Zoology 67:
198 – 207.
Silverman, H., Achberger, E. C., Lynn, J. W,. Dietz, T. H. 1995. Filtration and utilization of laboratory-cultured bacteria by Dreissena polymorpha, Corbicula fluminea and Carunculina texasensis. Biological Bulletin (Woods Hole, Mass.) 189:308 – 319.
Silverman, H., Lynn, J. W., Achberger, E. C., Dietz, T. H., 1996a. Gill
structure in zebra mussels: bacterial-sized particle filtration.
American Zoologist 36:373 – 384.
Silverman, H., Lynn, J. W., Achberger, E. C., Dietz, T. H. 1996b. Particle capture by the gills of Dreissena polymorpha: structure and
function of the latero-frontal cirri. Biol. Bull (Woods Hole,
Mass.) 191:42 – 54.
Silverman, H., Nichols, S. J., Cherry, J. S., Achberger, E. C., Lynn, J.
W., Dietz, T. H. 1997. Clearance of laboratory-cultured bacteria
by freshwater bivalves: differences between lentic and lotic
unionids. Canadian Journal of Zoology 75:1857 – 1866.
Silverman, H., Lynn, J. W., Beninger, P. G., Dietz, T. H. 1999. The
role of latrero-frontal cirri in particle capture by the gills of
Mytilus edulis. Biological Bulletin (Woods Hole, MA)
197:368 – 376.
Singh, D K., Thakur, P. K., Munshi, J. S. D. 1991. Food and feeding
habits of a freshwater bivalve, Parreysia favidens (Benson) from
the Kosi River System. Journal of Freshwater Biology 3:287 – 293.
Smith, B. J., Kershaw, R. C. 1979. Field guide to the non-marine molluscs of south eastern Australia. Australian National University
Press, Canaberra.
Smith, D. G. 1985a. A study of the distribution of freshwater mussels
(Mollusca: Pelecypoda: Unionidae) of the Lake Champlain
drainage in northwestern New England. American Midland
Naturalist 114:19 – 29.
11. Mollusca: Bivalvia
Smith, D. G. 1985b. Recent range expansion of the freshwater mussel, Anadonta implicata and its relationship to clupeid fish
restoration in the Connecticut River system. Freshwater Invertebrate Biology 4:105 – 108.
Smith, L. M., Vangilder, L. D., Hoppe, R. T., Morreale, S. J., Brisbin,
I. L., Jr. 1986. Effect of diving ducks on benthic food resources
during winter in South Carolina, U.S.A. Wildfowl 37:136 – 141.
Sparks, B. L., Strayer, D. L. 1998. Effects of low dissolved oxygen on
juvenile Elliptio complanata (Bivalvia: Unionidae). Journal of
the North American Benthological Society 17:129 – 134.
Sprung, M. 1991. Costs of reproduction: a study on the metabolic requirements of the gonads and fecundity of the bivalve Dreissena
polymorpha. Malacologia 33:63 – 70.
Sprung, M., Rose, U. 1988. Influence of food size and food quantity
on the feeding of the mussel Dreissena polymorpha. Oecologia
77:526 – 532.
Stanczykowska, A. 1984. Role of bivalves in the phosphorus and nitrogen budget in lakes. Verhandlungen-Internationale Vereinigung fur Theoretische und Angewandte Limnologie 22:982 – 985.
Stansbery, D. H. 1970. 2. Eastern freshwater mollusks. (I.) The Mississippi and St. Lawrence River systems. American Malacological Union Symposium on Rare and Endangered Mollusks.
Malacologia 10:9 – 22.
Stansbery, D. H. 1971. Rare and endangered freshwater mollusks in
eastern United States. in: Jorgensen, S. E., Sharp, R. E., Eds.
Proceedings of a symposium on rare and endangered mollusks
(naiads) of the U.S. Region 3, Bureau Sport Fisheries and
Wildlife, U.S. Fish Wildlife Service. Twin Cities, MN, pp.
5 – 18f.
Starrett, W. C. 1971. Mussels of the Illinois River. Illinois Natural
History Survey Bulletin 30:264 – 403.
Starnes, L. B., Bogan, A. E. 1988. The mussels (Mollusca: Unionidae)
of Tennessee. American Malacological Bulletin 6:9 – 37.
Stearns, S. C. 1980. A new view of life-history evolution. Oikos
35:266 – 281.
Steffens, W. L., Silverman, H., Dietz, T. H. 1985. Localization and
distribution of antigens related to calcium-rich deposits in the
gills of several freshwater bivalves. Canadian Journal of Zoology 63:348 – 354.
Stern, E. M. 1983. Depth distribution and density of freshwater mussels (Unionidae) collected with scuba from the lower Wisconsin
and St. Croix Rivers. Nautilus 97:36 – 42.
Stewart, T. W., Miner, J. G., Lowe, R. L. 1998. Quantifying mechanisms for zebra mussel effects on benthic macroinvertebrates:
organic matter production and shell-generated habitat. Journal
of the North American Benthological Society 17:81 – 94.
St. John, F. L. 1982. Crayfish and bivalve distribution in a valley in
southwestern Ohio. Ohio Journal of Science 82:242 – 246.
Stoeckel, J. A, Schneider, D. W., Soeken, L. A., Blodgett, K. D.,
Sparks, R. E. 1997. Larval dynamics of a riverine metapopulation: implications for zebra mussel recruitment, dispersal, and
control in a large river system. Journal of the North American
Benthological Society 16:586 – 601.
Stone, N. M., Earl, R., Hodgson, A., Mather, J. G., Parker, J., Woodward, F. R. 1982. The distributions of three sympatric mussel
species (Bivalvia: Unionidae) in Budworth Mere, Cheshire. Journal of Molluscan Studies 48:266 – 274.
Strayer, D. 1983. The effects of surface geology and stream size on
freshwater mussel (Bivalvia: Unionidae) distribution in southeastern Michigan, U.S.A. Freshwater Biology 13:253 – 264.
Strayer, D. L. 1993. Macrohabitats of freshwater mussels (Bivalvia:
Unionacea) in streams of the northern Atlantic Slope. Journal of
the North American Benthological Society 12:236 – 246.
Strayer, D. L. 1999a. Use of flow refuges by unionid mussels in
rivers. Journal of the North American Benthological Society
427
18:468 – 476.
Strayer, D. L. 1999b. Effects of alien species on freshwater mollusks
in North America. Journal of the North American Benthological Society 18:74 – 98.
Strayer, D. L., Jirka, K. J. 1997. The pearly mussels of New York
State. The New York State Museum Memoir 25, The University
of the State of New York, The State Education Department,
pp. xiii, 1 – 113.
Strayer, D. L., Ralley, J. 1993. Microhabitat use by assemblage of
stream-dewelling unionaceans (Bivalvia), including two rare
species of Alasmidonta. Journal of the North American Benthological Society 12:247 – 258.
Strayer, D. L., Caraco, N. F., Cole, J. J., Findlay, S., Pace, M. L.
1999. Transformation of freshwater ecosystems by bivalves.
Bioscience 49:19 – 27.
Summathi, V. P., Chetty, A. N. 1990. Ambient ammonia clearance by
bivalve mollusc Lamellidens corrianus (Lea). Environment and
Ecology 8:1333 – 1334.
Tankersley, R. A. 1996. Multipurpose gills: effect of larval brooding
on the feeding physiology of freshwater unionid mussels. Invertebrate Biology 11:243 – 255.
Tatem, H. E. 1986. Bioaccumulation of polychlorinated biphenyls
and metals from contaminated sediment by freshwater prawns,
Macrobrachium rosenbergii and clams, Corbicula fluminea.
Archives of Environmental Contamination and Toxicology
15:171 – 183.
Taylor, R. W. 1985. Comments on the distribution of freshwater
mussels (Unionacea) of the Potomac River headwaters in West
Virginia. Nautilus 99:84 – 87.
Taylor, R. W., Spurlock, B. D. 1982. The changing Ohio River naiad
fauna: a comparison of early Indian Middens with today. Nautilus 96:49 – 51.
Tevesz, M. J. S., Cornelius, D. W., Fisher, J. B. 1985. Life habits and
distribution of riverine Lampsilis radiata luteola (Mollusca: Bivalvia). Kirtlandia 41:27 – 34.
Thompson, C. M., Sparks, R. E. 1977. Improbability of dispersal of
adult Asiatic clams, Corbicula manilensis, via the intestinal tract
of migratory water fowl. American Midland Naturalist 98:
219 – 223.
Thompson, C. M., Sparks, R. E. 1978. Comparative nutritional
value of a native fingernail clam and the introduced Asiatic
clam. Journal of Wildlife Management 42:391 – 396.
Thorp, J. H., Bergey, E. A. 1981. Field experiments on responses of a
freshwater, benthic macroinvertebrate community to vertebrate
predators. Ecology 62:365 – 375.
Timm, H., Mutvei, H. 1993. Shell growth of the freshwater unionid
Unio crassus from Estonian rivers. Proceedings of the Estonian
Academy of Sciences Biology 42:55 – 67.
Trueman, E. R. 1983. Locomotion in molluscs. in: Saleuddin, A. S.
M., Wilbur, K. M., Eds. The Mollusca. Vol. 4: Physiology. Part
1. Academic Press, New York, pp. 155 – 198.
Turgeon, D. D., Bogan, A. E., Coan, E. V., Emerson, W. K., Lyons,
W. G., Pratt, W. L., Roper, C. F. E., Scheltema, A., Thompson,
F. G., Williams, J. D. 1988. Common and scientific names of
aquatic invertebrates from the United States and Canada: Mollusks. American Fisheries Society Special Publication 16,
Bethesda MD.
Turgeon, D. D., Quinn, J. F., Jr., Bogan, A. E., Coan, E. V.,
Hochberg, F. G., Lyons, W. G., Mikkelsen, P., Neves, R. J.,
Roper, C. F. E., Rosenberg, G., Roth, B., Scheltema, A.,
Sweeney, M. J., Thompson, F. G., Vecchione, M., Williams,
J. D. 1998. Common and scientific names of aquatic invertebrates from the United States and Canada: Mollusks, American
Fisheries Society Special Publication 26, 2nd ed., pp. 536 [Also
on CD-ROM].
428
Robert F. McMahon and Arthur E. Bogan
Ussary, T. A., McMahon, R. F. 1994. Comparative study of the dessication resistance of zebra mussels (Dreissena polymorpha) and
quagga mussels (Dreissena bugensis). in: Proceedings: Fourth
international zebra mussel conference ‘94. Wisconsin Sea Grant
Institute, Madison, WI, pp. 351–369.
van der Schalie, H. 1938. Contributing factors in the depletion of naiads in eastern United States. Basteria 3:51 – 57.
van den Thillart, G., de Vries, I. 1985. Excretion of volatile fatty
acids by anoxic Mytilus edulis and Anodonta cygnea. Comparative Biochemistry and Physiology B80:299 – 301.
Vaughn, C. C. 1997. Regional patterns of mussel species distributions in North American rivers. Ecography 20:107 – 115.
Vaughn, C. C., Pyron, M. 1995. Population ecology of the endangered Ouachita rock–pocketbook mussel, Arkansia wheeleri
(Bivalvia: Unionidae), in the Kiamichi River, Oklahoma. American Malacological Bulletin 11:145 – 151.
Vaughn, C. C., Taylor, C. M. 1999. Impoundments and the decline of
freshwater mussels: a case study of an extinction gradient. Conservation Biology 13:912 – 920.
Vaughn, C. C., Taylor, C. M. 2000. Macroecology of a host – parasite
relationship. Ecography 23:11 – 20.
Vaughn, C. C., Taylor, C. M., Eberhard, K. J. 1997. A comparison of
the effectivenes of timed searches vs. quadrat sampling in mussel surveys. in: Cummings, K. S., Buchanan, A. C., Mayer, C.
A., Naimo, T. J., Eds. Conservation and management of freshwater mussels II: initiatives for the future. Proceedings of a
Symposium, 1995, Upper Mississippi River Conservation Committee, Rock Island, IL, pp. 157 – 162.
Vermeij, G. J., Dudley, E. C. 1985. Distribution of adaptations: a
comparison between the functional shell morphology of freshwater and marine pelecypods. in: Trueman, E. R., Clarke, M.
R., Eds. The Mollusca. Vol. 10: Evolution. Academic Press,
New York, pp. 461 – 478.
Vidrine, M. F. 1990. Fresh-water mussel – mite and mussel –
Ablabesmyia associations in Village creek, Hardin County,
Texas. Proceedings of the Louisiana Academy of Sciences
53:1 – 4.
Vidrine, M. F. 1993. The historical distributions of freshwater mussels in Louisiana. Gail Q. Vidrine Collectibles. Eunice, LA.
Wade, W. C., Hudson, R. G., McKinney, A. D. 1993. Comparative
response of Ceriodaphnia dubia and juvenile Anodonta imbecillis to selected complex industrial whole effluents. in: Cummings,
K. S., Buchanan, A. C., Koch, L. M., Eds. Conservation and
management of freshwater mussels. Proceedings of a Symposium, 1992, Upper Mississippi River Conservation Committee,
Rock Island, IL, pp. 109 – 112.
Walker, B. 1918. The Mollusca. In: Ward, H. B., Whipple, G. C.,
Eds. Fresh-water Biology, New York, New York, pp.
957 – 1020.
Waller, D. L., Rach, J. J., Cope, W. G., Luoma, J. A. 1993. A sampling method for conducting relocation studies with freshwater
mussels. Journal of Freshwater Ecology 8:397 – 399.
Walz, N. 1978a. The energy balance of the freshwater mussel Dreissena polymorpha Pallas in laboratory experiments and in Lake
Constance. I.. Patterns of activity, feeding and assimilation efficiency. Archiv fur Hydrobiologie 55(Supp.):83 – 105.
Walz, N. 1978b. The energy balance of the freshwater mussel Dreissena polymorpha Pallas in laboratory experiments and in Lake
Constance. III. Growth under standard conditions. Archiv fur
Hydrobiologie 55(Supp.):121 – 141.
Warren, M. L., Jr., Cicerello, D. R., Camburn, K. E., Fallo, G. J.
1984. The longitudinal distribution of the freshwater mussels
(Unionidae) of Kinniconick Creek, northeastern Kentucky.
American Malacological Bulletin 3:47 – 53.
Watters, G. T. 1992. Unionids, fishes, and the species area curve.
Journal of Biogeography 19:481 – 490.
Watters, G. T. 1993. Mussel diversity as a function of drainage area
and fish diversity: management implications. in: Cummings, K.
S., Buchanan, A. C., Koch, L. M. Eds. Conservation and management of freshwater mussels. Proceedings of a Symposium,
1992, Upper Mississippi River Conservation Committee, Rock
Island, IL, pp. 113 – 116.
Watters, G. T. 1994a. Form and function of unionoidean shell scupture and shape (Bivalvia). American Malacological Bulletin
11:1 – 20.
Watters, G. T. 1994b. Annotated bibliography of the reproduction and
propagation of the unionoidea (primarily of North America).
Ohio Biological Survey Miscellaneous Contributions 1:1 – 159.
Watters, G. T. 1995. A guide to the freshwater mussels of Ohio. Division of Wildlife, The Ohio Department of Natural Resources.
Way, C. M. 1988. An analysis of life histories in freshwater bivalves
(Mollusca: Pisidiidae). Canadian Journal of Zoology 66:
1179 – 1183.
Way, C. M. 1989. Dynamics of filter-feeding in Musculium transversum (Bivalvia: Sphaeriidae). Journal of the North American
Benthological Society 8:243 – 249.
Way, C. M., Wissing, T. E. 1982. Environmental heterogeneity and
life history variability in the freshwater clams, Pisidium variable
(Prime) and Pisidium compressum (Prime) (Bivalvia: Pisidiidae).
Canadian Journal of Zoology 60:2841 – 2851.
Way, C. M., Wissing, T. E. 1984. Seasonal variability in the respiration of the freshwater clams, Pisidium variabile (Prime) and P.
compressum (Prime) (Bivalvia: Pisidiidae). Comparative Biochemistry and Physiology A78:453 – 457.
Way, C. M., Hornbach, D. J., Burky, A. J. 1980. Comparative life
history tactics of the sphaeriid clam, Masculium partumeium
(Say), from a permanent and temporary pond. American Midland Naturalist 104:319 – 327.
Way, C. M., Hornbach, D. J., Burky, A. J. 1981. Seasonal metabolism of the sphaeriid clam, Musculium partumeium, from a permanent and temporary pond. Nautilus 95:55 – 58.
Way, C. M., Hornbach, D. J., Deneka, T., Whitehead, R. A. 1989. A
description of the ultrastructure of the gills of freshwater bivalves, including a new structure, the frontal cirrus. Canadian
Journal of Zoology 67:357 – 362.
Way, C. M., Hornbach, D. J., Miller-Way, C. A., Payne, B. S., Miller, A.
C. 1990a. Dynamics of filter-feeding in Corbicula fluminea (Bivalvia: Corbiculidae). Canadian Journal of Zoology. 68:115 – 120.
Way, C. M., Miller, A. C., Payne, B. S. 1990b. The influence of physical factors on the distribution and abundance of freshwater
mussels (Bivalvia: Unionacea) in the lower Tennessee River.
Nautilus 103:96 – 98.
Weber, C. I. 1973. Biological field and laboratory methods for measuring the quality of surface waters and effluents (macroinvertebrate section). EPA-670 / 4-73-001. National Environment Research Center, Office of Research and Development, United
States Environment Protection Agency, Cincinnati, OH.
White, D. S. 1979. The effect of lake-level fluctuations on Corbicula
and other pelecypods in Lake Texoma, Texas and Oklahoma.
in: Britton, J. C., Ed. Proceedings, first international Corbicula
symposium. Texas Christian University Research Foundation,
Fort Worth, Tx, pp. 82 – 88.
Wilbur, K. M., Saleuddin, A. S. M. 1983. Shell formation. in: Saleuddin, A. S. M., Wilbur, K. M., Eds. The Mollusca. Vol. 4: Physiology, Part 1. Academic Press, New York, pp. 236 – 287.
Wilcox, S. J., Dietz, T. H. 1995. Potassium transport in the freshwater bivalve, Dreissena polymorpha. Journal of Experimental Biology 198:861 – 868.
11. Mollusca: Bivalvia
Wilcox, S. J., Dietz, T. H. 1998. Salinity tolerance of the freshwater
bivalve Dreissena polymorpha (Pallas, 1771) (Bivalvia: Dreissenidae). Nautilus 111:143 – 148.
Williams, C. J. 1985. The population biology and physiological ecology of Corbicula fluminea (Müller) in relation to downstream
dispersal and clam impingement on power station raw water
systems. Master’s Thesis, The University of Texas at Arlington,
Arlington, Tx.
Williams, C. J., McMahon, R. F. 1985. Seasonal variation in oxygen
consumption rates, nitrogen excretion rates and tissue organic
carbon: nitrogen ratios in the introduced Asian freshwater bivalve, Corbicula fluminea (Müller) (Lamellibranchia: Cobiculacea). American Malacological Bulletin 3:267 – 268.
Williams, C. J., McMahon, R. F. 1986. Power station entrainment of
Corbicula fluminea (Müller) in relation to population dynamics,
reproductive cycle and biotic and abiotic variables. American
Malacological Bulletin Special Edition No. 2:99 – 111.
Williams, C. J., McMahon, R. F. 1989. Annual variation of tissue
biomass and carbon and nitrogen content in the freshwater bivalve, Corbicula fluminea, relative to downstream dispersal.
Canadian Journal of Zoology 67:82 – 90.
Williams, J. D., Fradkin, A. 1999. Fusconaia apalachicola, a new
species of freshwater mussel (Bivalvia: Unionidae) from precolumbian archaeological sites in the Apalachicola Basin of Alabama, Florida, and Georgia. Tulane Studies in Zoology and
Botany 31:51 – 62]
Williams, J. D., Fuller, S. L. H., Grace, R. 1992. Effects of impoundments on freshwater mussels (Mollusca: Bivalvia: Unionidae) in
the main channel of the Black Warrior and Tombigbee Rivers in
western Alabama. Bulletin of the Alabama Museum of Natural
History 13:1 – 10.
Williams, J. D., Warren, M. L., Jr., Cummings, K. S., Harris, J. L.,
Neves, R. J. 1993. Conservation status of freshwater mussels of
the United States and Canada. Fisheries 18:6 – 22.
Willows, A. O., Ed. 1985. The Mollusca Vol. 8: Neurobiology and
behavior, Part 1. Academic Press, New York.
Willows, A. O., Ed. 1986. The Mollusca Vol. 9: Neurobiology and
behavior, Part 2. Academic Press, New York.
Winnell, M. H., Jude, D. J. 1982. Effect of heated discharge and entrainment on benthos in the vicinity of the J. H. Campbell
Plant, Eastern Lake Michigan, 1978 – 1981. Special Report No.
94. Great Lakes Research Division, University of Michigan,
Ann Arbor, MI, 202 pp.
Wissing, T. E., Hornbach, D. J., Smith, M. S., Way, C. M., Alexander, J. P. 1982. Caloric contents of corbiculacean clams (Bivalvia: Heterodonta) from freshwater habitats in the United
States and Canada. Journal of Molluscan Studies 48:80 – 83.
Woody, C. A., Holland-Bartels, L. 1993. Reproductive characteristics
of a population of the washboard mussel Megalonaias nervosa
429
(Rafinesque 1820) in the upper Mississippi River. Journal of
Freshwater Ecology 8:57 – 66.
Wu, W-L., Trueman. 1984. Observation on surface locomotion of
Sphaerium corneum (Linneaus) (Bivalvia: Sphariidae). Journal
of Molluscan Studies 50:125 – 128.
Wu, W-L. 1987. The burrowing of Anodonta cygnea (Linnaeus,
1758) (Bivalvia: Unionidae). Bulletin of Malacology Republic of
China 13:29 – 41.
Yamada, A., Matsushima, O. 1992. The relation of d-alanine and
alanine racemase activity in molluscs. Comparative Biochemistry and Physiology B103:617 – 621.
Yeager, M. M., Cherry, D. S., Neves, R. J. 1994. Feeding and burrowing behaviors of juvenile rainbow mussels, Villosa iris (Bivalvia: Unionidae). Journal of the North American Benthological Society 13:217 – 222.
Young, M. R., Williams, J. 1983a. The status and conservation of the
freshwater pearl mussel Margaritifera margaritifera Linn. in
Great Britain. Biological Conservation 25:35 – 52.
Young, M. R., Williams, J. 1983b. Redistribution and local recolonization by the freshwater pearl mussel Margaritifera margaritifera (L.). Journal of Conchology 31:225 – 234.
Young, M. R., Williams, J. C. 1983c. A quick secure way of making
freshwater pearl mussels. Journal of Conchology 31:190.
Young, M. R., Williams, J. 1984a. The reproductive biology of the
freshwater pearl mussel Margaritifera margaritifera (Linn.) in Scotland II. Laboratory studies. Achiv für Hydrobiologie 100: 29–43.
Young, M. R., Williams, J. 1984b. The reproductive biology of the
freshwater pearl mussel Margaritifera margaritifera (Linn.) in
Scotland I. Field studies. Archiv für Hydrobiologie 99: 405 – 422.
Zale, A. V., Neves, R. J. 1982. Fish hosts of four species of lampsiline
mussels (Mollusca: Unionidae) in Big Moccasin Creek, Virginia.
Canadian Journal of Zoology 60:2535 – 2542.
Zbeda, T. W., White, D. S. 1985. Ecology of the zoobenthos of
southeastern Lake Michigan near the D. C. Cook Nuclear
Plant. Part 4: Pisidiidae. Special Report No. 13. Great lakes Research Division, Institute of Science and Technology, University
of Michigan, Ann Arbor, MI.
Zeto, M. A., Tolin, W. A., Smith, J. E. 1987. The freshwater mussels
of the upper Ohio River, Greenup and Belleville Pools, West
Virginia. Nautilus 101:182 – 185.
Zheng, H., Dietz, T. H. 1998a. Paracellular solute uptake in the
freshwater bivalves Corbicula fluminea and Toxolasma texasensis. Biological Bulletin (Woods Hole, MA) 194:170 – 177.
Zheng, H., Dietz, T. H. 1998b. Ion transport in the freshwater bivalve Corbicula fluminea. Biological Bulletin (Woods Hole,
MA) 194:161 – 169.
Zs.-Nagy, I., Holwerda, D. A., Zandee, D. I. 1982. The cytosomal
energy production of molluscan tissues during anaerobiosis.
Basic and Applied Histochemistry 26 (Suppl.):8 – 10.
This Page Intentionally Left Blank
12
ANNELIDA: OLIGOCHAETA,
INCLUDING BRANCHIOBDELLIDAE
Ralph O. Brinkhurst
Stuart R. Gelder
205 Cameron Court
Hermitage, TN 37076
Department of Biology
University of Maine at Presque Isle
Presque Isle, ME 04769
OLIGOCHAETA
I. Introduction to the Oligochaeta
II. Oligochaete Anatomy and Physiology
A. Anatomy
B. Physiology
III. Ecology and Evolution of Oligochaeta
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Collecting, Rearing, and Preparation of
Oligochaetes for Identification
A. Collection
B. Rearing Oligochaetes
C. Preparation for Identification
V. Taxonomic Keys for Oligochaeta
A. Taxonomic Key to Families of
Freshwater Oligochaeta and
Aphanoneura
B. Taxonomic Key to Genera and
Selected Species of Freshwater
Lumbriculidae
C. Taxonomic Key to Genera and
Selected Species of Freshwater
Naididae
OLIGOCHAETA
I. INTRODUCTION TO THE OLIGOCHAETA
One family of aquatic Oligochaeta1, the Tubificidae (sludge worms), has been recognized since the time
of Aristotle for its ability to develop dense colonies in
1
Oligochaeta may be a parahyletic group. The term is used here in
the absence of a resolution into monophyletic taxa.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
D. Taxonomic Key to Genera and
Selected Species of Freshwater
Tubificidae
BRANCHIOBDELLIDAE
VI. Introduction to Branchiobdellidae
VII. Anatomy and Physiology of
Branchiobdellids
A. External Morphology
B. Organ-System Function
C. Environmental Physiology
VIII. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life history
C. Ecological Interactions
D. Evolutionary Relationships
IX. Identification of Branchiobdellids
A. Collecting, Rearing, and
Preparation for Identification
B. Taxonomic Key to Genera of
Freshwater Branchiobdellids
Literature Cited
Additional Suggested Readings
organically polluted waters (Fig. 1). Beginning with the
concepts that all sludge worms are pollution-tolerant
indicators, recent work has made biologists aware of
the wide range of ecological niches occupied by aquatic
annelids in general, and oligochaetes and branchiobdellids in particular. They are found in every kind of freshwater and estuarine habitat, not only in organic mud.
Specialized forms can be found in groundwater, in oligotrophic lakes and streams, and as commensals or
symbionts on crayfish, molluscs, and even tree frogs.
431
432
Ralph O. Brinkhurst, and Stuart R. Gelder
II. OLIGOCHAETE ANATOMY AND PHYSIOLOGY
A. Anatomy
FIGURE 1
Clump of living tubificids (photograph by P. M.
Chapman).
Several North American genera prey on other worms
or a variety of small invertebrates. The most surprising
recent discovery has been the description of a rich
diversity of tubificids at all depths in the oceans of
the world.
While most students are familiar with the terms
Oligochaeta and Clitellata, there is no general agreement about the equivalent rankings of the various annelid groups. Some authors place the arthropods and
annelids in a single phylum, but most texts separate
them. Within the Annelida, it is possible to classify the
Aclitellata (including Polychaeta) and the Clitellata as
superclasses or classes. The former arrangement allowed us to refer to the Polychaeta, Oligochaeta,
Hirudinea, and Branchiobdellida as classes, whereas
the latter would demote these to orders. Relationships
among the last three taxa, as well as the monotypic
Acanthobdellida, are presently under investigation using cladistic methods, which greatly assist the process
of determining sister-group relationships. Results of recent cladistic analyses provide evidence of a sistergroup relationship between lumbriculids and branchiobdellids, which share a unique arrangement of
genital organs. As a result, we have returned to an earlier classification of branchiobdellids as a taxon of family rank. The Aeolosomatidae are now recognized as a
non-clitellate taxon Aphanoneura.
Biologists who reject Hennigian methods may
well classify the various groups in a different way
from those who do not. The various terms will be
used here based on the concept of superclass Clitellata
with a series of classes, although we strongly suspect
that the ranking of the Acanthobdellida will shortly
be revised.
Oligochaete worms are bilaterally symmetrical,
segmented coelomates with four bundles of chaetae on
every segment except the first. In earthworms, the
chaetal “bundles” usually consist of two chaetae each;
but in aquatic species there are often several, and there
may even be more than a dozen per bundle. In a few
exceptional forms, the chaetae are absent in some or
even all segments. Aquatic species (superorder Microdrili) are smaller than earthworms (Megadrili) as the
name suggest, and they have simple body walls and
none of the specialized regions of the digestive tract
found in the earthworms. The clitellum is single-layer
thick, and is limited to the genital regions in microdriles, and the eggs are large and yolky. In earthworms,
secretions from the multilayered clitellum provide an
alternative source of food for the eggs, which are small
and less yolky. The clitellum is located well behind the
gonadal segments. In aquatic species, the male and female ducts are connected to external genital pores located either in the same segment as the gonads they
serve, or in the segment immediately behind them. In
earthworms, the male ducts are extended back to open
well behind the gonadal segments. It is generally
thought that there were originally four gonadal segments, two with paired testes, and two with paired
ovaries, in X – XIII (Roman numerals are used for segment number by convention). These have been reduced
in number in various ways, as gamete storage and
sperm transmission through copulation have evolved
(see Section III.D). In some entire families and in a few
individual species in other families, the gonadal segments may be found much nearer to the anterior end
than usual. This may be due to a tendency to regenerate fewer than normal number of anterior segments in
the predominantly asexual-reproducing forms in which
this happens (family Naididae). This explanation will
not suffice for sexually reproducing forms in which the
forward shift involves just one segment (family Lumbriculidae). In the family Opistocystidae, the gonads
are located behind segment XX in all but the one
species discussed here.
The arrangement of the various reproductive organs of some typical aquatic oligochaetes is illustrated
in Figure 2. The reproductive system and chaetae of
one side are illustrated, but in life these animals are bilaterally symmetrical. The anatomy of the reproductive
system is easy to understand if the stages in reproduction from sperm development to egg hatching are described. The mother cells of the sperm leave the testes
to float free in the coelom. The amount of space
12. Annelida: Oligochaeta, Including Branchiobdellidae
433
FIGURE 2 Reproductive systems and external characteristics of three major families of Oligochaeta. The
worms are shown with the anterior to the left, and only the structures of the left side are visible. The dorsal
and ventral chaetae are illustrated. In the Naididae, an example with dorsal chaetae beginning in VI is
shown. Segments are identified with Roman numerals by convention (septa would be VII/VIII, VIII/IX, etc.).
a, Atrium; e, eye spot; o, ovary; pr, prostomium; pt, prostate; sp, spermatheca; t, testis; v, vas deferens; genital pores.
available to the developing sperm masses, or morulae,
is increased by the development of outpushings of the
testicular segments, slightly anteriad but more extensively posteriad through several segments. These are
called sperm sacs. The sperm eventually cluster around
large funnels suspended on the posterior walls of the
testicular segment, sometimes well within the sperm
sacs. These sperm funnels lead into vasa deferentia that
originally drained via the male pores, located near the
ventral chaetae of the post-testicular segment. Various
structures have evolved, presumably at the point where
the vasa deferentia contact the original body wall. Inversion of the body wall has led to the evolution of
atria. These muscular bodies were originally lined with
ectodermal cells, many of which had a glandular function. They presumably fed and lubricated the sperm
stored in the atria ready for copulation. The glandular
cells are thought to have migrated inward, through the
muscle layers, so that their cell bodies now lie in the
coelom. This resulted in both an increase in the space
available to the glandular cell bodies and an increase in
the space available for sperm within the atria. The
434
Ralph O. Brinkhurst, and Stuart R. Gelder
ways in which the gland cells have migrated through
the muscles of the atria to form the prostate glands
vary, and this provides a very useful taxonomic feature
(though not one that can be seen without some effort).
The term prostate gland is used for very different structures in earthworms and branchiobdellids. In a further
evolutionary development, another section of the body
wall becomes part of the male reproductive structure
between the atria and the male pores. Various types of
penes are found in what are considered to be the more
advanced groups (family Tubificidae, family Lumbriculidae) and have evolved quite independently several
times. In some penes, the normal cuticular layer over
the epidermis has become hypertrophied to form cuticular penis sheaths. Sometimes this cuticular layer is
hardly noticeable, but the cuticular tubes may become
thick and rigid with soft penes free within them. Cuticular penis sheaths are said to be present if the cuticle
over the penes can be seen in cleared, whole-mounted
worms and has a consistent shape. In some species,
basement membranes or cuticular lining of eversible
penes appear on whole mounts, but these are usually
irregular in form. These are called “apparent penis
sheaths.”
When worms copulate, sperm is passed to the partner and is usually deposited in saclike spermathecae.
There may be one or more pairs of these structures located in or near the gonadal segments, commonly in
front of the testicular segments, or in them. The number and position of the spermathecae may vary within
the Lumbriculidae, but there is usually one pair in the
single testicular segment (families Tubificidae, and Naididae), or separated from the other genitalia in V (family Enchytraeidae). In a few species, spermathacae may
be present in some specimens but not others, or they
may be missing altogether. In Bothrioneurum (Tubificidae), the sperm are placed in spermatophores that are
attached externally to the body wall of the partner,
usually near the gonopores, during copulation. In
members of the tubificid subfamily Tubificinae, two
types of sperm are present, which come together in organized masses called spermatozeugmata. These can be
recognized within the spermathecae of mated worms.
Other forms of spermatozeugmata are found in other
subfamilies.
After a worm has copulated and has sperm in its
spermathecae, the clitellum secretes a cocoon. The eggs
have already developed in the coelomic space of the
ovarian segment. This segment is extended posteriad
into egg sacs. As the sperm sacs already displace the
septa of several segments behind the testes, the egg sacs
and their contained eggs are only detectable in segments beyond the sperm sacs. The egg and sperm sacs
have been omitted from Figure 2 because it becomes
confusing to follow a whole series of fingerlike protrusions running through several segments, all contained
within each other. [Note that egg and sperm sacs of the
Acanthobdellida and Hirudinea are not contained
within each other, but lie parallel to each other. They
are paired, bilateral structures just like the rest of the
reproductive system.] Eggs are deposited into the cocoon via the female ducts. These consist of nothing
more than ciliated funnels on the posterior walls of the
ovarian segment that open into the intersegmental furrow. In a few primitive forms, the female ducts actually
penetrate the postovarian segment and open on its anterior surface. It is assumed that fertilization occurs in
the cocoon. The basic position for the spermathecal,
male, and female pores is in the longitudinal line of the
ventral (ventrolateral) chaetal bundles, but there is
some variation in this due to midventral fusion, the development of median copulatory bursae, or the migration of the spermathecal pores to a more dorsal position. While the spermathecal pores may open near the
anterior furrow of the segment they occupy, all the genital pores commonly lie close to the ventral chaetae.
The worm finally releases the cocoon, which
formed as a ring of material around it, secreted (presumably) by the clitellum. The ends of the cocoon
close, and the fertilized eggs develop inside. Small
worms eventually escape through the openings at the
ends of the cocoons and are recognizable as immature
specimens, which can be identified if they happen to
belong to one of the species with characteristic chaetae.
The various reproductive organs and the modified
chaetae that may develop close to the genital pores can
best be found by looking carefully in the correct segments. When the keys mention genital structures, these
will only be found on mature specimens. An immature
aquatic oligochaete appears to consist of very little
more than the body wall with its chaetae and a simple
tubular gut running through the coelom. Very few specialized anatomical features have been added to the basic vermiform shape, unlike the situation in the Polychaeta. That is why it is generally useless to illustrate
the general habit of oligochaetes; useful illustrations
are those of the chaetae and reproductive systems. In
contrast to immature specimens, a mature worm will
have an external clitellum, though this is not as prominent as that of an earthworm, and the egg and sperm
sacs are full and, therefore, visible. The worm is often
somewhat distended in the genital region, and the reproductive organs obscure the normal view of the gut.
As most worms burrow through soft substrates
and extract their food by ingesting them, the range of
externally visible anatomical adaptations are few because this habit imposes severe limitations on body
shape. The prostomium lies in front of the mouth, and
12. Annelida: Oligochaeta, Including Branchiobdellidae
while its body cavity is clearly coelomic, it is considered to be presegmental. The prostomium may bear a
median anterior prolongation, the proboscis. In Bothrioneurum, there is a median dorsal pocket containing
sensory cells, the prostomial pit (Fig. 3). Segment numbering begins with the peristomium, the small segment
around the mouth. In aquatic species, the peristomium
is completely separate from the prostomium. It is also
presegmental, and bears no chaetae. This is important
to remember when using cleared, whole-mounted specimens as it is often easier to locate specific segments by
counting chaetal bundles (which begin in Segment II).
The chaetae vary in form as well as number. This
variation provides very useful key characters, but there
is increasing evidence that the fine details of chaetal
form, or even the presence or absence of a particular
type of chaetae, such as hairs, can be affected by quite
simple environmental variables such as pH or conductivity (Chapman and Brinkhurst, 1987). The Enchytraeidae have simple-pointed chaetae, which may
be straight or sigmoid. The chaetae in a bundle
may vary in size (Fig. 4). Hair chaetae are elongate,
simple shafts. They are found only in the dorsal bundles of some species of Naididae, Tubificidae, and in
Crustipellis (Opistocystidae). They are present in both
dorsal and ventral bundles in the Aeolosomatidae
(Aphanoneura, Aclitellata), this being one of a series
FIGURE 4
FIGURE 3
435
Prostomial pit, Bothrioneurum (photograph by P. M.
Chapman).
of fundamental differences between these minute
freshwater worms and true oligochaetes. Hair chaetae
are termed hispid or serrate when they bear a series of
fine lateral hairs. This condition may vary within a
Representative chaetae. From the left, top row: lumbriculid bifid with reduced upper tooth, head
end, and entire; small dorsal and large ventral sickle-shaped chaetae of Haplotaxis; three types of bundles from
the Enchytraeidae. Bottom row: two pectinates, a palmate, and a bifid dorsal chaeta, Tubificidae; six types of
dorsal needles, Naididae.
436
Ralph O. Brinkhurst, and Stuart R. Gelder
species depending on seasonal or environmental variables, but it is still used as a taxonomic character for
many naidids for which the limits of intraspecific variability have yet to be established. In a few naidids, the
lateral hairs may be especially prominent along one
side of the hair chaeta, and these are called plumose
hairs from their featherlike appearance. In one or two
instances, minute bifid tips have been observed at the
ectal end of hair chaetae, indicating their probable origin from the much more ubiquitous bifid chaetae. This
bifid form, with the ectal end divided into two teeth, is
characteristic of aquatic as opposed to terrestrial
oligochaetes, although the functional reason for this is
unknown. Bifid chaetae may be the only type present
in all bundles, and the ventral bundles usually contain
nothing else. Some simple-pointed chaetae may be
found in a few anterior ventral bundles (in the tubificid genus Spirosperma, for instance), or they may accompany the hair chaetae in the dorsal bundles in
some Naididae. A common combination in the dorsal
bundles is hair chaetae accompanied by pectinate
chaetae in, at least, the preclitellar bundles. Pectinate
chaetae are usually of the same general form as the bifids for the particular taxon, but the space between the
two teeth is filled with either a comb of thin teeth or,
more rarely, a ridged web. In a few naidids and tubificids, these webbed chaetae may be expanded as
palmate chaetae. In general, the dorsal chaetae accompanying the hair chaetae in the majority of naidid
species differ markedly from the ventral chaetae. For
this reason, they have come to be termed needles,
whether they are palmate, pectinate, bifid, or simplepointed. Some authors like to substitute the terms fascicle, capilliform, bifurcate, and crotchet for bundle,
hair, bifid, and chaeta, respectively. They also use the
terms distal and proximal for upper and lower tooth,
the designations used in describing the variations in
the length and thickness of the teeth, often a useful
characteristic. [Anglo-Saxon terms are more succinct,
and are used internationally, but the longer Latin
terms are more readily translated into French and
Spanish.] The normal ventral chaetae may be replaced
by specialized genital chaetae, usually beside the spermathecal or penial pores. These are illustrated where
appropriate in the keys. Genital chaetae do not vary
much in form in the Naididae, where there are often
two or three blunt penial chaetae in each bundle beside the male pore. These chaetae are quite long proximally, but the distal end beyond the nodulus (the
swelling on the chaeta at the point where it emerges
from the chaetal sac) is short. Penial chaetae are usually somewhat long and thicker than the normal ventral chaetae. Penial chaetae, like those of the Naididae,
may be found in the Rhyacodrilinae, a subfamily of
Tubificidae, though there are usually several to a bundle. They are usually arranged with the outer ends
close together, the inner ends spread out. In Tubificinae, the ventral chaetae in the penial segment are usually lost but the spermathecal chaetae are often modified, with elaborate distal ends shaped like trowels.
They may be used to introduce the modified spermatozeugmata into the spermathecae during copulation.
The functions of both these and the penial chaetae as
claspers are based on speculation because aquatic
oligochaetes have rarely been observed mating. Spermathecal chaetae are often associated with special
glands. Where these are large, and there is a great deal
of competition for space in the genital segments, the
spermathecal chaetae and glands lie immediately in
front of the spermathecal segment (Rhizodrilus, Tubificidae). In general, the number and diversity of chaetae
is reduced toward the posterior end of the worm, and
modified teeth on the bifids commonly become “normal.” There are a few exceptions. The upper teeth of
the chaetae of Amphichaeta (Naididae) get longer posteriorly, and the posterior segments of Telmatodrilus
vejdovskyi (Tubificidae) bear chaetae with almost
brushlike tips (see illustrations in the key).
A few other anatomical features are used in
aquatic oligochaete taxonomy. External gills are found
in a few taxa. The posterior segments of the tubificid
Branchiura bear single dorsal and ventral gill filaments. These are longest in the median segments of the
gill-bearing region, very short at each end of the row.
Dero (Naididae) has gills at the posterior end of the
body, usually enclosed in a gill chamber (branchial
fossa). These are best observed in living specimens. All
aquatic oligochaetes have red blood pigments which
aid oxygen uptake and transport. In most species, respiration takes place through the posterior body wall or
even within the pre-anal part of the intestine. Water
may be pumped in and out of the anus for this purpose. Most aquatic oligochaetes protrude the tail end
out of the sediment into the water and may wave the
tail to create enough disturbance to maintain high concentrations of oxygen around the tail. The body wall
of posterior segments in many lumbriculids is penetrated by special branching blood vessels, presumably
to improve respiration.
External sense organs are mostly limited to ultrastructural features of the body wall (Jamieson, 1981).
These may become prominent in those species that protect the body wall with secretions that trap bacteria
and foreign matter in an external crust. Sensory papillae can be seen protruding through such layers in living
specimens, but they are usually retracted in fixed material. This has led to arguments about their presence or
absence in certain species, but we may assume that any
12. Annelida: Oligochaeta, Including Branchiobdellidae
worm with its normal body-wall sensory system denied
access to information by being encrusted has developed
localized retractile sensory papillae. It is also easy to
understand why the necessarily sensitive prostomium
and first two or three segments may be naked in taxa
that have protected body walls, and why the anterior
end is often retractile within the rest of the protected
body. Simple pigmented eye spots are located on the
lateral surface of segment I in a number of naidids, but
may be present or absent in various specimens of the
same species.
The majority of aquatic oligochaetes feed by ingesting sediment, extracting nutrition from the organic
matter and especially the bacteria, which comprise less
than 10% of it. They feed continuously in a conveyorbelt fashion, often ingesting recolonized feces. Food is
obtained by everting the roof of the pharynx through
the mouth. Glandular cells associated with the pharynx
have evolved into pharyngeal or septal glands, in which
the cell bodies lie in the coelomic cavities of several anterior segments, attached to the posterior septa. Intercellular canals run forward from these cell bodies along
the lateral sides of the gut to open dorsally on the pharyngeal pad, which is one of the characteristics of
worms of this group; this feature is absent in the Aeolosomatidae. The pharyngeal glands are used in taxonomy of the Enchytraeidae, but not in other families.
The pharyngeal glands are lost in predatory
oligochaetes, but traces of the dorsal pad can still be
found in some (Chaetogaster, Naididae) although it has
been lost in others (Phagodrilus, Lumbriculidae, and
Haplotaxis, Haplotaxidae).
B. Physiology
The primary emphasis of studies on worm physiology has been on respiration, as it provides a simple
model of the value of red blood pigment function and
anaerobic metabolism. A good review of respiration
studies with reference to their use in determining production in worms was provided by Bagheri and
McLusky (1984), who studied the estuarine tubificid
Tubificoides benedii (benedeni). Oxygen consumption
varied from 0.01 to 0.5 L / mg / h, but while there
were small, apparent linear increases in rate with temperature, these were not statistically significant. These
results are among the lowest reported for worms, suggesting that the experimental technique is not creating
stress in the test animals. The response of respiration
levels to stress has been suggested as a tool in pollution
biology (Brinkhurst et al., 1983). Most tubificids
respire at a more or less constant rate as the external
oxygen concentration is lowered. Once a critical value
is reached, respiration rate falls dramatically. Stress can
437
elevate or depress the constant respiration rate or it
may shift the critical value. It may also cause a reduction in, or a total loss of, respiratory control above the
critical value. In some instances, respiratory control actually increased in the face of supposed stress factors. A
unique phenomenon is that mixing certain tubificid
species together will actually reduce their respiration
rates by a significant amount (Chapman et al., 1982a).
This result was anticipated by earlier studies of worm
production using monocultures and mixed cultures of
three tubificid species. In those studies with mixed populations (summarized in Brinkhurst and Austin, 1978),
growth rates increased when respiration decreased, as
did assimilation efficiencies. These experiments seemed
to provide an explanation for the way in which tubificids live in small clumps of mixed species. Some worms
preferred the recolonized feces of another species as
food and sought the company of that species. Even
when they were only in water from monocultures of
the appropriate second species, worms moved less, ate
more, and (in the presence of living worms) ingested
more of their preferred food. This caused a shift in all
of the variables measured in production work based on
estimating caloric or carbon budgets. The work provided an unusual example of the value of multi-species
flocks.
Freshwater oligochaetes, such as the pollutiontolerant tubificids Tubifex tubifex and Limnodrilus
hoffmeisteri, can withstand exposure to 10 ppt salinity,
but others can only survive at 5 ppt (Chapman et al.,
1982b, c). A low level of salinity actually seems to improve the ability of freshwater worms to withstand
stress factors. It is less surprising that the provision of
sediment improves stress resistance. Estuarine species
may withstand a very wide range of environmental
stress, including extreme salinity values (Paranais, Naididae; Monopylephorus, Tubificidae). It would be a
mistake to believe that these animals are exposed to a
wide range of salinities due to daily tidal excursions,
however. In muddy situations in rivers such as the
Fraser in British Columbia, the “interstitial” salinity
goes through a seasonal cycle related to the capture
and release of snow in the headwaters. As temperatures
increase, the flow of freshwater increases and tidal seawater incursions are limited to the outer estuary. All of
the benthic communities move slowly up and down the
estuary in response to this annual rhythm of interstitial
salinity, maintaining themselves at what are presumably preferred salinity ranges for each species. It is selection for these optimal salinities (that remain quite
constant within the mud on short-term basis) that
causes the sequence of species observed along the salinity gradient in estuaries (Chapman, 1981; Chapman
and Brinkhurst, 1981).
438
Ralph O. Brinkhurst, and Stuart R. Gelder
While worms in cocoons may be able to survive
desiccation, there is no experimental evidence of this.
Adult Tubifex tubifex (Tubificidae) are able to form
cysts for adult worms that can survive unmoistened
for 14 days, or for as long as 70 days with occasional
dampening. Once the worms have been moistened,
they may emerge in 20 h (Kaster and Bushnell, 1981).
The lumbriculid Lumbriculus variegatus can also encyst, but it only survived for 4 days under experimental conditions. In both instances, the covering of foreign matter, which helps to disguise the cyst, is also
essential to the survival of the worms. There are
records of a handful of other lumbriculids undergoing
fission in cysts, but no physiological work has been
done. Lumbriculus also forms a different type of cyst
to withstand freezing (Olsson, 1981). Specimens of the
tubificid Limnodrilus profundicola were collected by
R. Brinkhurst at a height of ca. 4000 m on Mount
Evans, Colorado. After being accidentally frozen into
a solid block of ice, the worms emerged quite unscathed when thawed the next day. As so many
aquatic oligochaetes, especially the Naididae, are cosmopolitan or at least very widely distributed, one
would expect more evidence of adaptations to counter
desiccation. Dispersal mechanisms of oligochaetes are
still a matter for speculation.
The development of eye spots is unique to the Naididae, a family of small, delicate worms, some of which
can make rudimentary swimming movements. Other
naidid species, while not free-living, occupy tubes attached to aquatic plants and may protrude the anterior
end. The majority of aquatic oligochaetes avoid the
light by keeping the head end buried in their food supply, the sediment. Most aquatic worms have quite sensitive tails and have a rapid withdrawal reflex based on
a mechanosensory detection system that is responsive
to the approach of potential predators (Zoran and
Drewes, 1987). Branchiura has the fastest escape reaction reported for any invertebrate. This speed is associated with the development of the lateral giant nerve
fibers in the ventral nerve cord. These are responsible
for rapid forward transmission of sensory information.
The median giant nerve fiber transmits posteriad from
the head end. In Lumbriculus, which lives in shallow
littoral habitats, the body is extended up to the
air – water interface, with the tail lying horizontally in
it. Not surprisingly, these worms have what Drewes
and Fourtner (1988) termed hindsight or light-sensitive
tails, and they respond to passing shadows. Light can
be damaging to worms.
A great deal of work has been done in the laboratory on the effect of toxic substances on worms, summarized by Chapman and Brinkhurst (1984) and
Wiederholm et al. (1987).
III. ECOLOGY AND EVOLUTION OF OLIGOCHAETA
A. Diversity and Distribution
Many aquatic oligochaetes are cosmopolitan, or at
least widely distributed. This is especially true of the
Naididae, but even here significant patterns are beginning to emerge. The supposedly primitive naidid subfamily Stylarinae is scarce in the southern United
States, such genera as Stylaria, Arcteonais, Ripistes,
and Vejdovskyella being common in northern locations. Dero and Allonais species (members of the Naidinae) are well represented in subtropical and tropical
regions. There are other north – south distribution patterns. The tubificid Arctodrilus wulikensis is found in
Alaska, as are several lumbriculid species that are part
of the Pacific Rim fauna mentioned later. Phallodrilus
hallae, a tubificid limited to Lake Superior, is related to
a large number of marine taxa in the subfamily Phallodrilinae. Unlike the northern forms, tubificid species restricted to southern or southeastern localities belong to
genera that are otherwise more widely distributed.
Among the tubificids and lumbriculids, some taxa
are limited to the western United States and Canada.
Some of these are related to or are the same as Asian
Pacific Rim counterparts, suggesting an evolutionary
distribution pattern. Several tubificids are known only
from Lake Tahoe, which may prove to be home to a
number of endemic species, though some forms first described there have since been found elsewhere, mostly
still in California.
The apparent limitation of some species to the
eastern United States may simply reflect a greater frequency of caves or more interest in cave faunae in
those areas. At least six lumbriculid species fall into
this category, but there are also some eastern surface
water tubificid species. The groundwater tubificid Rhyacodrilus falciformis is a European species now known
to be widespread in such habitats on Vancouver Island,
British Columbia, and in Illinois, and there is a lumbriculid known from a well in California; so there may be
as rich a western groundwater fauna (awaiting description) as there is in eastern North America. Many
groundwater species have very limited distributions, although the distinctions between species in genera such
as Trichodrilus are often based on minute differences.
This may mean that biologists look more carefully for
differences where they anticipate that they may exist.
The reasons for these geographic patterns have not
been studied, partly because they have only been recognized for a few years. There may be ecological reasons
for the distribution of some taxa. Species of Aulodrilus
(Tubificidae) may be widely distributed, but they
appear to reproduce asexually in the more temperate
12. Annelida: Oligochaeta, Including Branchiobdellidae
zones . This may suggest a recent spread from warmer
areas where they do breed sexually. Some species can
be shown to be recent introductions by humans. The
North American Limnodrilus cervix has been introduced to Britain and Sweden via ports and canals (Milbrink, 1983). It is possible that the diverse collection of
Potamothrix (Tubificidae) species in the St.
Lawrence – Great Lakes represents a series of introductions of common European species into North America. This is more certain for Psammoryctides barbatus
(Tubificidae), a very recent discovery in the St.
Lawrence River in Quebec (Vincent et al., 1978), as
well as the estuarine and coastal species Tubificoides
benedii found recently by the author in Vancouver
Harbor, British Columbia. One of the most common
tropical tubificids, Branchiura sowerbyi, was first described from a Victorian greenhouse in Britain, where
it had been introduced along with the tropical plants
on display. It has now become adapted to temperate
waters, though it still thrives best in reservoirs and
canals, especially where the temperature is elevated. It
has apparently spread all across North America since it
was first recorded in Ohio in 1931; but, this could simply reflect a gradual spread of awareness of this easily
recognized species (Brinkhurst, 1965). Some of these
regional distributions are mentioned in the keys, but
the reader should always be prepared for the unusual.
Niche discrimination in aquatic oligochaetes is less
obvious than zoogeographic patterns. The majority of
these worms are adapted to live in sediments ranging
from sand to mud. They can be found in pockets of
such sediments in stony habitats as well as in lowland
rivers, lakes, and ponds where soft substrates are the
norm. There is none of the usual distinction between
lacustrine and riverine species. The best guess that can
be made about niche discrimination is that the precise
nature of the organic matter in sediments, and the
microflora it supports, determines the outcome of
interspecific competition. The ecophysiological work
on multispecific flocks described earlier is probably
applicable to most situations and not limited to
the Tubifex – Limnodrilus – Quistadrilus association in
which it was initially described. Milbrink (1993) obtained similar results with T. tubifex and Potamothrix
moldaviensis. In most habitats, between 5 and 15
species can be found. There are some special adaptations, such as the groundwater fauna already mentioned. Several genera of the Naididae are no longer restricted to life in the sediment, but may be found
among aquatic weeds in quiet rivers and ponds. Lumbriculus lives among vegetation in pond and stream
margins. Dero (Allodero) species are symbionts of
frogs, which, once released from their host come to resemble specimens of Dero (Aulophorus), rendering the
439
taxonomic distinction suspect. The predators Haplotaxis and Phagodrilus are found in springs, seepages,
and groundwater, but Chaetogaster is more ubiquitous.
Estuarine species form another group, but a number of
species are restricted to tidal freshwater where there is
little, if any, detectable saltwater. None of these exhibit
any marked anatomical adaptations to their way of
life, with the exception of the pharyngeal structures of
the predators and loss of chaetae and gills in Allodero.
The significance of food in the ecological specialization of the aquatic oligochaetes, especially the tubificids, is demonstrated by the sequence of species groups
that inhabit progressively more organically polluted
stretches of rivers or more eutrophic lakes. The Lumbriculidae and some naidid genera such as Pristina and
Pristinella are found in streams with low organic matter in the sediments, and they may be accompanied by
tubificids belonging to the Rhyacodrilinae in particular
(Rhyacodrilus, Bothrioneurum, and Rhizodrilus). Tubificinae, such as Potamothrix and Aulodrilus, can be
found in situations with a reasonable supply of organic
matter but also a good oxygen supply, although
Tubifex tubifex, several Limnodrilus species, and Quistadrilus are found in areas where the oxygen supply
can become reduced. This basic pattern has been expanded, quantified, and recognized in Europe as well
as North America (e.g., Lang, 1984). At first, this type
of work followed the much earlier pattern of establishing lists of chironomid midge species characteristic of
lake types (reviewed in Brinkhurst, 1974). The maximum value in identifying aquatic oligochaetes in pollution studies today is to add information for use in data
matrices to be analyzed by multivariate statistics, such
as cluster analyses. There are relatively few invertebrate
species in freshwater muddy habitats, in contrast to
stony streams (where there are far more insects) or marine habitats (where there are many more muddwelling taxa). The taxonomic identites of the communities from each sample site are retained in such
analyses, whereas they are lost in simpler diversity
models.
B. Reproduction and Life History
Field studies on reproduction in the Naididae were
reviewed by Loden (1981). Asexual reproduction, (usually by paratomy), predominates, but sexual reproduction apparently enables populations to persist through
periods of unsuitable conditions. Abundance varies seasonally depending on the geographic region, with
spring, summer, or winter maxima being produced
asexually. Most species apparently cease feeding as
they mature because the gut degenerates, and the
mouth may even become sealed. According to Loden,
440
Ralph O. Brinkhurst, and Stuart R. Gelder
some species incorporate the anterior part of the worm
in the cocoon, but Lochhead and Learner (1984) disputed this in describing a life history for Nais. They reported that worms from this and other naidid genera
are able to shed their cocoons. According to these investigators, the adult worms are more resistant to environmental stress than the cocoons. All cocoons of
aquatic oligochaetes may be disguised with mud particles, or may be placed out of reach of predators (Newrlka and Mutayoba, 1987).
Most tubificids reproduce sexually, but there are
several parthenogenetic species. The latter often lack
spermathecae and the testes may also be reduced or absent. The vasa deferentia may be reduced to solid
strands in these species, but the atria are usually retained. A few other species seem able to reproduce
asexually by fission. The results of various field studies
produce conflicting interpretations of the life histories
of such common genera as Limnodrilus. There is less
disagreement about events in less adaptable species
found in more restricted habitats. The worms may mature a few weeks after hatching if temperature and
food supplies are adequate, and they may breed more
than once in a season. Alternative studies suggest that
many species mature in their second year. After breeding, these forms resorb the gonads and gonoducts and
would be recognized as immature specimens. The next
year they develop a new set of reproductive organs and
breed again, after which most species die. Some published examples are the studies of Block et al. (1982)
and Lazim and Learner (1986a, b) for Limnodrilus
species, Christensen (1984) on asexual reproduction,
and Poddubnaya (1984) on parthenogenesis. Because
of the problems encountered in interpreting the results
of field studies on life histories, considerable effort has
been put into the development of laboratory cohort
studies (Bonacina et al., 1987). In general, oligochaete
life histories are probably not as precisely tuned to environmental variables like day length as they are in
more highly evolved forms such as insects. This may be
attributable to simpler neurosecretory systems in
worms, but there is no evidence to support this idea.
Food and temperature may cause considerable local
variation in life cycles, especially in the opportunistic
species like Limnodrilus hoffmeisteri. Unfortunately,
these abundant, cosmopolitan species are the most convenient laboratory animals, but they are so adaptable
and perhaps atypical that comparable work should be
done with species found in more restricted habitats.
C. Ecological Interactions
There is some published work on the food and
feeding behavior of the Naididae, but a much larger
literature exists on the relationship between tubificids
(and one lubriculid species Stylodrilus heringianus) and
the sediments that they ingest (and therefore defecate)
continuously.
Algae and other epiphytic material are the main
food of many naidids, but Nais feeds on heterotrophic,
aerobic bacteria like the tubificids. There is no specialized gut flora in either group (Brinkhurst and Chua,
1969; Harper et al., 1981). There is no evidence of selective ingestion or digestion of specific bacteria in
Nais, but selective digestion is important in tubificids.
Nais elinguis will select algal filaments in preference to
monofilament nylon thread and prefers some algae to
others or to glass beads (Bowker et al., 1985). Algae
also form part of the food supply of the tubificids
Limnodrilus claparedeianus and Rhyacodrilus sodalis
and the lumbriculid Lumbriculus variegatus.
Studies of defecation rate and the chemical and
bacterial content of feces in tubificids have usually exploited the fact that these worms will continue to feed
with their heads in the sediment and their tails in the
water column, even when such colonies in small containers are inverted. If a layer of cotton or fine netting
is used to prevent the sediment drifting down past the
worms into the collecting funnel placed beneath the
culture, it is easy to determine the amount and the
chemical content of the feces. If the feces are collected
over very short time periods, the bacterial flora of the
feces can be studied. Similar analyses of the sediment
provided as food make it possible to study the effect of
passage through the gut on the organic matter content
and bacterial flora (Brinkhurst and Austin, 1978). Fecal samples can also be collected from cultures oriented
normally, but the feces then have to be collected from
the artificial sediment surface instead of being allowed
to drop into a container. Both methods are potentially
liable to error and cultures must be handled very carefully. The behavior of the worms is affected by changes
in temperature and light and by physical disturbance.
The results derived from cultures maintained in both
orientations have usually shown that worms are not affected by being inverted, and that, if either method is
subject to error in determining the amount of fecal material produced, these errors cancel out. Only Kaster et
al. (1984) suggested that worms in the normal position
produce more feces (45 – 110%) than those in inverted
cultures. Such obvious differences would have been detectable in earlier studies had they existed. Defecation
rate seems to be very difficult to determine. The nutritional value of the food provided may decline as the experiment proceeds, and this may cause an increase in
the amount of sediment processed in order to feed the
colony. Experimental results are affected by periods of
acclimation to the artificial sediments, which may be as
12. Annelida: Oligochaeta, Including Branchiobdellidae
long as a month. Two supposedly identical cultures
maintained side by side may behave quite differently
(Appleby and Brinkhurst, 1970). Defecation rates may
be used in studies of worm production when associated
with estimates of organic matter content and caloric
value of the food and feces. They may also be used in
studies of sedimentation and bioperturbation of sedimentary layers in freshwater ecosystems. The lumbriculid Stylodrilus heringianus builds up large populations in many regions of the Great Lakes, where it
behaves in the same way as a tubificid. Krezoski et al.
(1984) used gamma spectroscopy methods to measure
the rates at which a layer of sediment (labeled with radioactive cesium) and overlying water (labeled with radioactive sodium) were transported by the burrowing
activity of worms and other benthos. The results of
these and similar studies all support the idea that the
superficial sediment layers in lakes are thoroughly
mixed by benthic organisms, and that solute transport
across the mud – water interface is greatly enhanced by
them. Worms are probably the most important animals
involved in this because of the depths to which they
penetrate the sediment, the large populations that they
build up, and their ceaseless activity. Gardner et al.
(1981) showed that invertebrates could be responsible
for most of the phosphorus released from aerobic sediments in Lake Michigan. Chatarpaul et al. (1979)
showed that worms accelerated nitrogen loss from
coarse stream sediments, indicating their importance in
rivers as well as lakes. There is now a growing realization that the sediments are not just the final resting
place for organic matter and contaminants, but that
they may become a source for them (Karlckhoff and
Morris, 1985).
D. Evolutionary Relationships
There are contradictory ideas about the evolution
of any biological group, many of which are due to fundamental differences in the methods used. There are
strongly held opinions about the way in which relationships should be determined, even among those who
employ cladistic methods and those who agree with at
least the basic concepts of Hennig. The following account is a personal one, based on experience with most
families of aquatic oligochaetes, an exposure to marine
and freshwater worms worldwide, and some familiarity
with cladistics. This is not said in order to validate this
particular view, but to help the reader identify the biases it inevitably includes. At least it should not suffer
from the geographic or taxonomic parochialism that
tends to cause taxonomic inflation — the tendency to
elevate the taxa in which we are most interested to the
highest level possible.
441
We should perhaps consider what the original
worm looked like, rather than asking whether polychaetes or oligochaetes came first. It seems quite reasonable to suppose that an animal looking a lot like a very
simple earthworm, but with a reproductive system that
released eggs and sperm freely into the environment,
provides a common ancestor to both groups (Fig. 5).
There has been some support for this idea among polychaete biologists, but, in general, they seem to have difficulty deciding on relationships above the family level,
and on what constitutes a primitive worm. The “simplified earthworm” with two chaetae per bundle and no
parapodia is adopted following the ideas of Clark
(1964) regarding the evolution of segments and coelom.
There is, of course, no reason to suppose that every
group evolved in the ocean. The teleost fish provide a
good example of a group that may well have undergone
a very large diversification in freshwater before re-invading the ocean, so that the existence of a large number of marine Tubificidae is not prima facie evidence for
their antiquity, even within that family, though the possibility should not be excluded. All possible combinations of the reproductive system segmental arrangement
within the oligochaetes (and other Clitellata) can be derived from a worm with four genital segments, two
with paired testes in each, followed by two with paired
ovaries. The original position of this series, termed GIGIV (Brinkhurst, 1982), seems to have been in X – XIII.
The organisms with this arrangement that exit today
are found in the Haplotaxidae, with only one or two
exceptions that are thought to be primitive members of
groups descended directly from that family (Fig. 6).
There are some exceptions that have been explained
elsewhere, but only the barest outline will be presented
here. [Challenges to this largely Austral group as being
ancestral come from the fact that Haplotaxis itself is a
highly modified genus of predatory worms (see the taxonomic key).] It is then thought that a number of families have quite independently lost the anterior testes (GI)
and the posterior ovaries (GIV), making copulation easier (there are fewer pores to bring into contact with
each other). This happens because the development of
egg and sperm sacs enables more gametes to be stored
in the coelom from a single pair of gonads than could
be achieved in a single segment when the sacs were
small or undeveloped. Similarly, the evolution of multiple chaetae per bundle seems to have arisen independently a number of times, most notably in the Enchytraeidae, Tubificidae, and Naididae. In the former, most
species have two chaetae per bundle, a few “specialists”
have no chaetae in some or all bundles (another common but independent evolutionary step found in several
families), but some have several chaetae. These are all
simple-pointed, however, which seems to be correlated
442
Ralph O. Brinkhurst, and Stuart R. Gelder
FIGURE 5
Phylogenetic tree of the Annelida. This diagram represents an outline of the major (potential)
monophylies and the apomorphic character states that define them (solid bars across lines). Note that the Branchiobdellidae is shown as included within the Oligochaeta, which may also be true for the Hirudinea according
to recent molecular evidence. Aphanoneura is expected to be eliminated as the Aeolosomatidae become placed
within the Polychaeta in future publication.
Phylogenetic tree of the Oligochaeta. While the Hirudinea (Acanthobdellida Euhirudinea) are
said to be related to the Lumbriculida according to molecular analyses, they do not have semiprosoporous
male ducts (possibly plesioporous and prosoporus respectively, a single pair of male ducts in Acanthobdella,
testes within sperm sacs confluent with the male pores with no male funnels in Euhirudinea). The form of the
male ducts in Acanthobdella and Euhirudinea could have arisen independently from a semiprosoporous condition, in which case they could form a monophyly or monophylies within the Lumbriculida, but the molecular
and anatomical schemes need to be resolved and hence the Hirudinea are excluded from this figure.
FIGURE 6
12. Annelida: Oligochaeta, Including Branchiobdellidae
with the fact that most members of this family are at
best only semiaquatic if not totally terrestrial. Chaetae
with bifid tips are a hallmark of aquatic oligochaetes, as
are more complex pectinate and palmate chaetae and
their derivatives, hair chaetae. The hair chaetae of the
Phreodrilidae (a small, Southern Hemisphere family)
seem to have evolved independently of those of the Naididae and Tubificidae. These are the two most closely
related families of aquatic oligochaetes, and they are apparently the most highly evolved in many respects. The
Naididae exhibit several adaptations to an aquatic life
as opposed to one restricted to burrowing in moist sediments. They have simple eyes, a few have specialized
chaetae in a few anterior segments, and some can swim
in a very simple way. The predominance of asexual reproduction enables them to exploit a more changeable
environment than that available in the sediments, and in
some as yet unidentified way has enabled most of them
to become very widely distributed. Their diet now includes algae as well as bacteria, and we can assume that
they can absorb macromolecules through the body wall
as food (as do the freshwater and marine tubificids).
The male reproductive system has become progressively
more complex in these groups. In the Enchytraeidae,
the male ducts may terminate in what appear to be
more glandular than muscular structures, which are
not yet identifiable as atria. The tubificids have muscular atria and the higher forms have stalked prostate
glands, elaborate penes, and either penis sheaths, penial
chaetae, or spermathecal chaetae.
The Lumbriculidae are more of a puzzle. They
have usually retained both testicular segments, but
have lost the second ovaries (GIV). Many advanced
forms secondarily loose GI, and a few parthenogenetic
forms have an irregular, asymmetrical, or even exaggerated number of gonads. The chaetae are restricted to
two per bundle, but these are usually bifid rather than
simple-pointed as postulated in the plesiomorphic condition. Some biologists see the paired chaetae as evidence of a close relationship between the lumbriculids
and the earthworms, suggesting that the latter are the
derivatives of these aquatic forms. This puts an enormous weight on just one character, but these people
also regard hair chaetae as being retained from a polychaete ancestor. The difficulty with relating the lumbriculids to the earthworms is that the lumbriculids (and
branchiobdellidans) have a unique way of reducing the
number of male pores. They have their atria in GII and
both pairs of male ducts drain into them. In Lumbriculids, the prostate glands consist of a diffuse covering
of gland cells and none of them have developed stalked
prostates. Cuticular penis sheaths are very rare; genital
chaetae are limited to a single species. Too much has
been made of the apparent diversity of reproductive
443
segments in this family. While the variation is greater
than that of other families, the most common form
does seem to be the ancestral one (Brinkhurst, 1989).
While the aquatic oligochaetes closest to their presumed ancestors seem to be found in Australia and
New Zealand, not unlike many other freshwater invertebrate groups, other families are widely distributed.
According to a cladistic analysis of some Enchytraeidae
(Coates, 1989), South America seemed to contain the
basal forms. In the Naididae, it is the northern Stylarinae that appear to the oldest (Nemec and Brinkhurst,
1987). The Lumbriculidae inhabit the Northern Hemisphere, with a very high proportion of them being limited to, or extending eastward from, Lake Baikal in
Siberia. The European Cookidrilus seems to most resemble the presumed ancestor, with the widespread
genera Stylodrilus and Trichodrilus being quite generalized (Brinkhurst, 1989). There has yet to be a detailed
analysis of the more cosmopolitan Tubificidae, but
there is a greater diversity of genera in the Northern
Hemisphere, with very few restricted to the Southern
Hemisphere (as Antipodrilus) and very few adapted to
the subtropical and tropical regions, apart from
Branchiura and, perhaps, Aulodrilus.
We are currently trying to assess the relationships
between the clitellate taxa usually termed Oligochaeta,
Hirudinea, and Acanthobdellida. In a preliminary revision (Brinkhurst and Gelder, 1989), it was shown that
the supposed link between the predatory lumbriculid
worm Agriodrilus from Lake Baikal (Russia) and the
non oligochaete taxa in this group is purely circumstantial, made even clearer by the recent discovery of a
parallel evolution in the North American Phagodrilus.
Without presenting all of the data here, it now looks as
if the branchiobdellids and Lumbriculidae share a common ancestor, both having two pairs of male ducts
draining through atria in GII, unless this character
proves to be a convergence. Hence, the former taxon is
referred to as Branchiobdellidae in this chapter. It remains to be seen if the male ducts of the Hirudinea and
the Acanthobdellida are at all related to what we might
call the semiprosopore state of the male ducts in the
lumbriculids (only one pair of male ducts are now
thought to be in their testicular segment in the ancestor
of the family according to a cladistic analysis). The extreme modification of the male ducts in the leeches, in
which the testes lie in sacs behind the two successive
segments bearing male and female pores, is considered
to be derived from a system with the testes in front of
the ovaries, as in the oligochaetes. With the loss of the
coelom, the testes are supposed to be transferred into
the surviving sperm sacs as a series of small gonads
replacing a single original pair. Complex male ducts
have been developed, but these are in the form of a
444
Ralph O. Brinkhurst, and Stuart R. Gelder
continuous tube in which the sperm are shed internally.
These ducts need to be reexamined for any trace of
relict sperm funnels, to see if there is any evidence that
they were ever prosoporous like the lumbriculids. The
acanthobdellids, represented by one species with one
probable synonym, have four pairs of chaetae in a few
anterior segments including the oral segment. There is
no trace of a prostomium, or peristomium, and perhaps some anterior segments (Brinkhurst, 1999). Acanthobdella also retains or possibly recovers the coelom
(which is lost in leeches), but the testes still lie within
the sperm ducts. Two intriguing aspects are the pattern
of eyes and the unique sperm-receiving device shared
by this genus and the piscicolid leeches. Other revised
information about the branchiobdellids, discussed later,
also weakens the argument for a relationship between
them and the leech groups. That argument was based
on a very superficial examination of the detailed facts
available. Textbook accounts in this and many other
areas of supposed evolutionary relationships should be
read with caution, especially if they give no clear indication of the theoretical basis on which the opinions
are based.
IV. COLLECTING, REARING, AND PREPARATION
OF OLIGOCHAETES FOR IDENTIFICATION
A. Collection
Oligochaetes may be collected using any of the
standard array of samplers from dip nets to grabs and
core tubes. Collectors specifically seeking worms can
use refined histological fixatives in the field. A great deal
of material is collected in the course of ecological surveys where time does not permit such care. Strictly
quantitative samples from soft sediments are best obtained with narrow core tubes without core retainers.
Using core tube lengths appropriate to the type of substrate can save a great deal of frustration, but most people seem to try to work with a single set of tubes. Soft
sediments can be washed through screens and sorted in
the laboratory, preferable after being preserved in their
entirety in the field. If cores are used, the volume to be
carried back from the field may be quite small. Field
sorting is relatively inaccurate, and small specimens
may well be missed. The best preservative for bulk samples is 10% buffered formalin (4% formaldehyde solution), but is easier to carry 40% in the field and add this
to an aqueous sample to approximate 10% in the
whole sample. Addition of rose bengal or phloxine B in
very small amounts aids sorting, and the stain can be
washed out of the worms when they are transferred to
70% alcohol for storage. The use of larger amounts of
stain creates problems in identifying worms. Sorting
should be done with stereomicroscopes under good
lighting. Care should be taken to avoid diluting the alcohol when transferring worms into the storage vial.
Many North American collections are sent for identification with the worms half decomposed because bulk
samples were preserved in alcohol, not formalin, or the
original alcohol has not been drained and replaced with
70%. Storage can be quite permanent in patent lip vials
with neoprene stoppers. The stoppers tend to swell and
seal the vial against alcohol loss. We have some in our
laboratories that have not needed attention for 20
years. If these vials and stoppers are not available, vials
must be sealed with tape, wax, or parafilm. Some neoprene stoppers dry and crack with age, and all collections should be curated to maintain their integrity. Museum quality material should be donated to institutions
that can provide curation.
If the collector is not taking quantitative samples
for ecological or environmental studies and is working
in a remote or unusual location, then it may be worth
the effort of sorting, fixing and preserving some specimens while in the field if they are readily visible. Putting
the sample in an enamel dish and allowing everything
to settle will reveal even small worms as they begin to
move. Carrying whole samples back to the laboratory
to sort them while the worms are alive can cause selective mortalities, as some species surviving elevated temperatures and lowered oxygen concentration better than
others (and the latter always seem to be the more interesting species from a taxonomic viewpoint). Live
worms can be sorted from sediment and detritus by
spreading clean sand over the sample or putting the
sample on a screen set over clean water. The worms will
then actively migrate into the sand or water. This can be
useful in samples full of plant fragments.
Samples taken with dip nets or other methods that
include coarse substrates can be processed in the field
to reduce the volume and weight of the sample. Repeated, careful elutriation into a screen, screen bucket,
or back through the original dip net can be used, imitating a gold-panning technique. The amount of vigor
used can be selected in order to separate enough of the
fine fraction including all the specimens from the heavy
material. The coarse residues should, of course, be
carefully checked for large, heavy, or attached organisms, or specimens trapped in empty shells and other
unusual situations.
B. Rearing Oligochaetes
Culture of worms can be as simple as placing the
substrate and the worms in an aerated aquarium,
preferable where temperatures can be kept at appropri-
12. Annelida: Oligochaeta, Including Branchiobdellidae
ate levels, usually between 4 – 10°C depending on the
original habitat. Small cultures used for laboratory experiments can be maintained in shallow vessels half
filled with sterilized lake sand. Filtered lake water is
added every two weeks and frozen lettuce strips are
buried in the sand to provide bacteria as food. Mass
culture to provide fish food can utilize manure to feed
the worms. Marian and Pandian (1984) described raising 7.5-mg Tubifex tubifex every 42 days on a mixture
of 75% cow dung and 25% fine sand, maintaining the
oxygen level at 3 mg / L. Fresh cow dung is added every
four days, and harvesting is supposedly best done at
night. Aston (1984) cultured Branchiura sowerbyi at
25°C. Leitz (1987) reviewed attempts to mass culture
Lumbriculus variegatus. Production rates of 15 kg / m2
and population doubling times of 11 – 42 days are reported. The use of live worms as fish food has many
advantages over other diets. Most commercially available live worms are now Lumbriculus (called ‘black
worms” in commerce) rather than Limnodrilus.
C. Preparation for Identification
Many biologists believe that these animals are hard
to identify. It is worth indicating that, as they are hermaphrodites, there is no difficulty with keys for males
and females, and there are no larval forms. About 60%
of the species can be identified from superficial somatic
characters, but the rest require mature specimens. It is,
of course, always preferable to confirm identities from
mature specimens wherever possible, but it is not essential. Those used to identifying organisms with stereomicroscopes may be bothered by the need to make slide
mounts, but simple methods can be used in most instances. The serious taxonomist will want to make
more elaborate preparations and may want to section
specimens to check details, but this is not necessary for
routine identification. There are only about 150 freshwater oligochaetes and a small number of branchiobdellidans in North America, and many of these are rare
and local. Recent keys have been written with the beginner in mind, and so they emphasize external or
readily visible internal characters. Such keys work well
if they proceed directly to the specific level, but it is a
little more difficult to write keys to the genera, as preferred in this volume. Generic characteristics are mostly
detailed aspects of the soft parts of the male reproductive structures. These may be difficult to see on rapidly
made preparations in which the soft parts are dissolved. The keys presented here do not always proceed
directly to genera for this reason, but we have managed
to write keys involving externally visible characteristics. Some reference to geographic distribution is made
in the keys, but this has been done only where the
445
information available seems reliable. Fully illustrated
keys with supporting descriptions of all the species are
available elsewhere (Kathman and Brinkhurst, 1998).
Once the worms are preserved in alcohol, they
must be prepared for study with a compound microscope. Examination of specimens from remote or unusual locations should begin with a study of the external characteristics, such as position of the gonopores
and other obvious features. However, for routine identifications of specimens from ordinary localities, very
little benefit can be derived from this step in the
process. In this case, the worms can either be immersed
in a temporary mounting medium by replacing the alcohol with Amman’s lactophenol, or they may be
washed with water and then placed in separate drops
of lactophenol on slides. It is usually possible to mount
live worms under a single coverslip (with two coverslips per slide) unless the worms are very large. These
wet mounts should be set aside for 1 – 2 days on stacking trays. Amman’s medium is quite corrosive, so it
should be handled with care and any spillage should be
wiped off microscope stages immediately. It consists of
20% phenol, 20% lactic acid, 20% water, and 40%
glycerine. It is best stored in dark bottles; however, it is
hygroscopic and does not have a good shelf life. Somewhat more permanent mounts can be made with media
such as CMCP and polyvinyl lactophenol. Such slides
should usually be sealed with a ring of material
(Glyceel, nail polish, etc.) around the edges of the coverslips to prevent the media from drying up.
None of these methods produce satisfactory permanent slides, nor are they adequate for working with
the Enchytraeidae or the marine Tubificidae, in which
internal anatomy must be seen. For this type of material, permanent whole mounts using stained and dehydrated worms are mounted in Canada Balsam or one
of its more recent substitutes. To prevent worms from
becoming brittle, they should be cleared in methyl salicylate rather than xylene. This will enable advanced
students to try bisecting (sagitally) the separated genital
region of each worm or even dissecting out the male
ducts when necessary. This should be limited to identification of new taxa to the generic level, when details
of the male reproductive system must be established.
Serial sections (usually sagital longitudinal) are also
employed in taxonomic work, but not in routine identification of well-known species. It is important to be
knowledgeable about the expected locations of the various organs before attempting dissection or interpretation of serial sections.
The procedure for examining a worm on a slide is
as follows. Check the prostomium for a proboscis or
sense organ. Next, examine the ventral chaetae of the
first two or three bundles and determine the form of the
446
Ralph O. Brinkhurst, and Stuart R. Gelder
dorsal bundles and where they begin (usually in II or
between IV and VI). Establish the number and form of
these chaetae (bifid, pectinate, hair chaetae, etc.). Determine the relative lengths of the teeth. As these are illustrated by taxonomists with the ectal or outer end of the
whole chaeta toward the top edge or corner of the figure, the upper tooth is always drawn lying above the
lower tooth. When viewing a whole-mounted specimen,
the chaetae often lie flat on the body wall and so no distinction of position can be made; therefore, always visualize a chaeta in the upright position. Care should be
taken to examine several chaetae from an exactly lateral
aspect, because slight deviations can produce apparent
distortion of the relative lengths of the teeth. Measurements often betray the bias of the eye of the observer
and are recommended where an eyepiece micrometer
scale is available. The worm should then be searched
for genital characters. Carefully check the appropriate
segments (usually X – XI in a tubificid, for example) to
see if the ventral chaetae are modified. If the dorsal and
ventral chaetae are alike, make sure that you have
examined at least three bundles in those segments to
ensure that a ventral bundle has been examined. Check
the penial segment to see if there is a penis sheath (they
should be paired, of course) even if it is thin and inconspicuous. Check the posterior chaetae and the body itself for gills or any other special features. Then proceed
to check the key. If the worm has distinct features, [like
a single dorsal hair with single simple needles (these beginning in V) and the anterior ventral (II – V) and posterior ventral (VI onward) bundles have chaetae that differ in length, width, and shape], it will rapidly become
possible to skip the key altogether and proceed straight
to Nais or Dero depending on the presence of gills.
Many such easy identifications to major groups or to
genera can be made with a little practice. Identifications
will require a properly aligned microscope, as chaetae
and penis sheaths have refractive indices that do not differ much from the background, and too much light
makes them impossible to see. Oil immersion lenses
must be used to see pectinations on chaetae in tubificids
(although with practice they can be identified without),
and must also be used to determine the form of the needles (dorsal chaetae) in naidids.
V. TAXONOMIC KEYS FOR OLIGOCHAETA
A. Taxonomic Key to Families of Freshwater Oligochaeta and Aphanoneura
1a.
Minute worms, 1 – 2 mm or chains of animals up to 10 mm; hair chaetae in both dorsal and ventral bundles; worms move in a gliding motion using cilia; body wall with colored or refractile epidermal glands; no eversible pharyngeal pad; no clitellum, only ventral
copulatory glands in rare mature specimens; nervous system ladderlike (Fig. 7)..............................................................Aphanoneura
.........................................................................................Aeolosomatidae [Key to world species in Brinkhurst and Jamieson (1971)]
1b.
Larger worms; hair chaetae restricted to dorsal bundles or absent; worms move by alternate contraction of longitudinal and circular
muscles; no colored epidermal glands; eversible pharyngeal roof present (except in aquatic earthworms); clitellum present in mature
worms; nervous system fused ventrally ..........................................................................................................Oligochaeta .................2
2a(1b).
Ventral chaetae large, sickle-shaped, single (i.e., two per segment) dorsal chaetae small, straight, often missing from some or all segments; prostomium very long, furrowed, no proboscis; mouth large, muscular pharynx present; body elongate, slender, resembling a
gordian worm (Fig. 8)........................................................................................................................Haplotaxidae............Haplotaxis
[Worms in this genus resemble the European H. gordioides (Hartman, 1821), but mature specimens have yet to be described from
North America. There may be several species (now regarded as synonyms) differing by the number and distribution of dorsal
chaetae.]
2b.
All chaetae paired, or multiple in a bundle, unless partially or totally absent; prostomium short, often conical, without a transverse
furrow, but with or without a proboscis; body form very rarely elongate and slender ........................................................................3
3a(2b).
Chaetae all paired from II or, if more numerous, all simple-pointed ...................................................................................................4
3b.
Chaetae more than two per bundle, usually bifid, sometimes with pectinate and hair chaetae dorsally, simple-pointed chaetae rare
(limited to a few anterior ventral bundles, or single needles with the hairs dorsally)...........................................................................7
4a(3a).
Chaetae paired, bifid, with small to rudimentary upper teeth (Fig. 9..........................Lumbriculidae (in part)..................(Section V.B)
4b.
Chaetae simple-pointed ......................................................................................................................................................................5
5a(4b).
Thick-bodied worms with clitellum several segments behind the gonopores (XII – XIV) .......................................................................
.......................................................................................................................Megadrili [Mostly terrestrial, but some aquatic species.
Most aquatic species are members of the Sparganophilidae (Sparganophilus) or the Lumbricidae (Eiseniella).]
5b.
Thin-bodied worms with clitellum one cell layer thick and in region of gonopores (X – XII or further forward) ................................6
12. Annelida: Oligochaeta, Including Branchiobdellidae
447
FIGURE 7
Aeolosoma sp. Note ciliated prostomium, lack of septa, simple pharynx, dorsal and ventral hair
chaetae, and colored body wall inclusions (irregular circles). FIGURE 8 Haplotaxis sp. Whole specimens can
be even longer in relation to the breadth. FIGURE 9 Bifid chaetae, Lumbriculidae. FIGURE 10 Enchytraeid
chaetae. These may be straight, curved, with or without noduale, and may differ in length within a bundle. All
chaetae are usually identical. The anterior end of Barbidrilus (below) have ventral bundles of strange rodlike
chaetae with bifid tips only in II and III, shown on the right.
6a(5b).
Larger worms (difficult to mount under a coverslip) with sigmoid, nodulate chaetae; proboscis present or absent; spermathecae in
or adjacent to the gonadal segments, male pores between VIII and X................................Lumbriculidae (in part) ..........(Section V.B)
6b.
Smaller worms with chaetae often straight, commonly not nodulate, sometimes missing from some if not all segments (in Barbidrilus Loden and Locy, 1981 with uniquely forked chaetae in II – III ventrally, others missing). No proboscis. Spermathecae open in
V, male pores in XII (Figs. 2 and 10) ..............................................................................................................................Enchytraeidae
7a(3b).
Worms (1.7 – 3.0 mm) with posterior end bearing one median and two lateral processes. Genital region with testes in XI, ovaries
and atria in XII, spermathecae in XIII (Fig. 11) .............................................................................................................Opistocystidae
[The only definite North American record is for Crustipellis tribranchiata (Harman, 1970), restricted to Louisiana, Mississippi,
Florida, and North Carolina. The tail processes make this obvious (Harman and Loden, 1978).]
7b.
Posterior end naked, or with gills, or with two lateral processes plus gills, no median process (Fig. 26); spermathecae in the testicular segment, atria in the ovarian segment, these being in X – XI or further forward (Fig. 2)................................................................8
8a(7b).
Length usually 1 cm; hair chaetae usually present in dorsal bundles but absent in some genera; hair chaetae commonly associated
with simple-pointed or bifid chaetae rather than the rarer palmate or pectinate chaetae, these dorsal chaetae (needles) very often differ from the ventrals in form (Fig. 12); dorsal chaetae may begin behind II, often in V or VI, sometimes elsewhere; dorsal chaetae
often limited to one or two needles, and one or two hairs per bundle; ventral bundles with numerous bifid chaetae, those of II or
even II – V often differ in form and thickness from the rest; asexual reproduction by budding forms chains of individuals; mature
specimens have spermathecae in IV, V or VI with the male pores one segment behind them (Fig. 2); eye spots may be present; gills
may surround the anus (Fig. 26); prostomium may bear a proboscis (Fig. 23.......................................Naididae .............(Section V.C)
8b.
Length usually 1 cm, width usually 0.5 – 1.0 mm; when hair chaetae are present dorsally, they are usually accompanied by pectinate chaetae that closely resemble the ventral chaetae, apart from having intermediate teeth (Fig. 13); when hair chaetae are absent
dorsally, all bundles usually contain similar bifid chaetae; dorsal chaetae begin in II, normally several per bundle; reproduction
448
Ralph O. Brinkhurst, and Stuart R. Gelder
FIGURE 11
Crustipellis (Opistocystidae). Tail end to left, head end with proboscis to right, genital segments
below (a, atrium, sp, spermatheca, both of left side). FIGURE 12 Various forms of dorsal chaetae, needles,
of Naididae. These usually differ from the more normally bifid ventrals. FIGURE 13 Dorsal chaetae of
Tubificidae. Pectinates, on the left, differ relatively little from a bifid, shown on the right. Ventral bundles
usually consist of bifids, sometimes with minute pectinations in those species with pectinate dorsals. Pectinates
are usually accompanied by hairs in the dorsal bundles, rarely are bifid dorsals accompanied by hairs. All
chaetae may be bifid.
normally sexual, rarely by fragmentation; spermathecal pores normally on X, male pores on XI (Fig. 2); no eyes or proboscis; no
gills around anus, single dorsal and ventral gill filaments on some posterior segments of one species (Fig. 30)....................Tubificidae
........................................................................................................................................................................................(Section V.D)
B. Taxonomic Key to Genera and Selected Species of Freshwater Lumbriculidae
1a.
Prostomium with a proboscis (Figs. 14 and 15)..................................................................................................................................2
1b.
Prostomium without a proboscis ........................................................................................................................................................6
2a(1a).
Chaetae bifid, at least in anterior segments.........................................................................................................................................3
2b.
Chaetae all simple-pointed .................................................................................................................................................................4
3a(2a).
Proboscis elongate; chaetae bifid in anterior segments (Fig. 14) ..................................................Kincaidiana hexatheca Altman, 1936
[This species is limited to the Pacific Northwest.]
3b.
Proboscis short; all chaetae bifid, but with colateral teeth (set side by side) (Fig. 15)............................................................................
Rhynchelmis brooksi Holmquist, 1976 .......................................................................................[Known only from northern Alaska.]
4a(2b).
Cave-dwelling species with short proboscis .............................................................................Trichodrilus allegheniensis Cook, 1971
[Known only from Tennessee.]
4b.
Surface water species with long proboscis ..........................................................................................................................................5
5a(4b).
Atria of male reproductive system with spiral muscles (Fig. 16) ...........................................................................Eclipidrilus (in part)
[This genus was reviewed by Brinkhurst (1988). The species with probosces are known mostly from the southeastern United States,
with one record from Montana.]
5b.
Atria without spiral muscles ...............................................................................................................................Rhynchelmis (in part)
[Several species distributed from California – Nevada to Alaska and Northwest Territories of Canada. See Fend and Brinkhurst
(2000)]
6a(1b).
Chaetae bifid ......................................................................................................................................................................................7
6b.
Chaetae simple-pointed ......................................................................................................................................................................8
7a(6a).
Elongate (to 100 mm or more) slender worms, the front end often green in life, the rest dark red to black; elaborately branched
blood vessels laterally in the body wall of posterior segments (Fig. 17); reproduces sexually and asexually (fragmentation with or
without encystment); no permanent everted penes............................................................................................................Lumbriculus
[One widely distributed species, Lumbriculus variegatus (Mller, 1774), inhabits shallow water, where it floats the tail in the
air – water interface. Several other possible taxa, mostly from Alaska, are almost certainly variants of this (see Lumbriculus and
Thinodrilus, in Holmquist 1976).]
7b.
Short (25 – 40 mm), tapering worms, pale to white color; blood vessels in the posterior lateral body wall with short, unbranched
12. Annelida: Oligochaeta, Including Branchiobdellidae
449
lateral diverticulae; reproduction sexual; permanently everted soft penes on X close together near the midline on mature specimens ..
..............................................................................................................................................Stylodrilus heringianus Claparéde, 1862
[Widespread in the Great Lakes and Canada, possibly a European introduction.]
8a.(6b).
Groundwater species ..........................................................................................................................................................................9
8b.
Surface water species ........................................................................................................................................................................11
9a(8a).
Four pairs of male pores on X, two pairs of spermathecal pores on IX (Fig. 18) ..........................Spelaedrilus multiporus Cook, 1975
[Known only from a cave in Virginia.]
9b.
Male pores paired on X, spermathecal pores paired in either IX, XI, or XI and XII .........................................................................10
10a.(9b).
Spermathecal pores paired on IX .........................................................................................................................................Stylodrilus
[S. californianus Rodriguez, 1996, California; S. wahkeenensis Rodriguez and Coates, 1996, Oregon, and possibly Tennessee and
Alabama.]
10b.
Spermathecal pores paired in XI or XI and XII....................................................................................................Trichodrilus (in part)
[From caves in West Virginia, Illinois, Washington.]
FIGURE 14 Proboscis and chaetae of Kincaidiana hexatheca. FIGURE 15 Proboscis of Rhynchelmis
brooksi. FIGURE 16 Spiral atrial muscles, Eclipidrilus. FIGURE 17 Blood vessels of the lateral body wall
of posterior segments, Lumbriculus. FIGURE 18 Ventral view of segments IX (to left) and X showing unique
gonopores of Spelaedrilus (sp, spermathecal pores; male symbol, male pores, 4 and 8 of each, respectively.)
FIGURE 19 Reproductive systems of Styloscolex, anterior to left, left side shown. Note testis (t) and atrium
in VIII, ovary (o) and female funnel in X, and spermatheca (sp) in XI. FIGURE 20 Reproductive system,
Kincaidiana freidris, anterior to left, organs of left side shown. Note testis (t) and atrium (a) in VIII, prostate
gland stalks shown as well as male duct, ovary (o), and spermatheca (sp) in IX.
450
Ralph O. Brinkhurst, and Stuart R. Gelder
11a(8b).
Male pores paired in VIII .................................................................................................................................................................12
11b.
Male pores paired in X.....................................................................................................................................................................13
12a(11a).
Spermathecal pores in XI, ovaries in X (Fig. 19) .............................................................Styloscolex opisthothecus Sokol’skaya, 1969
[This mostly Asian species is restricted to northern Alaska.]
12b.
Spermathecal pores paired in IX, ovaries in IX (Fig. 20) ....................................................................Kincaidiana freidris Cook, 1966
[This species may not be congeneric with K. hexatheca, but cladistic analyses suggest they are quite closely related. Both are Pacific
Northwest species, this one from California.]
13a(11b).
Spermathecal pores paired in IX ........................................................................................Stylodrilus sovaliki (Holmquist, 1976) and
.........................................................................................................................Tenagodrilus musculus Eckroth and Brinkhurst, 1996
[S. sovaliki is from Alaska, but it closely resembles its European congener S. absoloni (Hrabe, 1970). T. musculus is from Alabama.]
13b.
Spermathecal pores either single median in VIII – IX or in IX, or paired in IX .....................................................Eclipidrilus (in part)
[One species, (E. frigidus Eisen, 1881) is from Idaho and California; the other three (E. lacustris Verrill, 1871, E. ithys Brinkhurst,
1998, and E. fontanus Wassell, 1984) are from northeastern North America].
C. Taxonomic Key to Genera and Selected Species of Freshwater Naididae
1a.
No dorsal chaetae present (in some rare specimens the dorsal chaetae are recovered, but these can be recognized because of the absence of ventral chaetae in III – V and the presence of an enlarged pharynx and reduced prostomium associated with a predatory
habit) ...............................................................................................................................................................................Chaetogaster
1b.
Dorsal chaetae present (rare in Ophidonais) ......................................................................................................................................2
2a(1b).
Dorsal chaetal bundles without hair chaetae ......................................................................................................................................3
2b.
Dorsal chaetal bundles with hair chaetae (Fig. 2) ...............................................................................................................................7
3a(2a).
Dorsal chaetae begin in II or III ..........................................................................................................................................................4
3b.
Dorsal chaetae begin in V or VI .........................................................................................................................................................5
4a(3a).
Dorsal chaetae begin in II; no gap between chaetal bundles of III and IV....................................Homochaeta naidina Bretscher, 1896
4b.
Dorsal chaetae begin in III; gap between chaetal bundles of III and IV (Fig. 21) ..............................................................Amphichaeta
5a(3b).
Dorsal chaetae begin in V; estuarine species ............................................................................................................................Paranais
5b.
Dorsal chaetae begin in VI; freshwater species ...................................................................................................................................6
6a(5b).
Dorsal chaetae stout, solitary, bluntly simple-pointed or notched at the outer end (Fig. 22) ......Ophidonais serpentina (Müller, 1773)
6b.
Dorsal chaetae curved, with bifid tips, 2 – 4 per bundle. [See also Piguetiella, in which hair chaetae may be missing in most, if not
all, dorsal bundles; some Uncinais specimens are said to have dorsal chaetae in V.] .........................Uncinais uncinata (Orsted, 1842)
7a(2b).
With a proboscis on the prostomium (visible as a stump if broken off) (Fig. 23) ................................................................................8
7b.
Without a proboscis on the prostomium ..........................................................................................................................................11
8a(7a).
Dorsal chaetae begin in II .........................................................................................................................................................Pristina
8b.
Dorsal chaetae begin in VI (observe with care as ventral chaetae of IV and / or V may be missing).....................................................9
9a(8b).
Dorsal bundles of VI – VIII with 2 – 16 giant hair chaetae (Fig. 24) ...................................................Ripistes parasita (Schmidt, 1874)
9b.
No giant hair chaetae .......................................................................................................................................................................10
10a(9b).
Ventral chaetae with a characteristic double bend (Fig. 25); dorsal bundles with 1 – 3 hairs and 3 – 4 shorter, hairlike needles .............
........................................................................................................................................................Stylaria lacustris (Linnaeus, 1767)
10b.
Ventral chaetae slightly curved; dorsal bundles with 8 – 18 hairs and 9 – 12 hairlike needles .........Arcteonais lomondi (Martin, 1907)
11a(7b).
Dorsal chaetae begin in II or III ........................................................................................................................................................12
11b.
Dorsal chaetae begin in IV or further back .......................................................................................................................................14
12a(11a).
Body wall covered with foreign matter adhering to glandular secretions .....................................................................Stephensoniana
12b.
Body wall naked...............................................................................................................................................................................13
13a(12b).
Dorsal chaetae begin in II; hair chaetae in all bundles, none especially thin, either none especially elongate “or” elongate hair
chaetae in III .........................................................................................................................................................................Pristinella
12. Annelida: Oligochaeta, Including Branchiobdellidae
451
FIGURE 21
Anterior end (to right), Amphichaeta, showing dorsal chaetae beginning in III, and gap between
chaetae of III and IV, particularly pronounced in this species, but noticeable throughout the genus. FIGURE 22
Dorsal chaetae, Ophidonais. These are single, or even absent in many segments. FIGURE 23 Proboscis
on the prostomium of a naidid, this one with dorsal chaetae from II, but this varies (see key). FIGURE 24
Dorsal chaetae of Ripistes begin in VI, with giant hairs in three segments. FIGURE 25 Ventral chaeta of
Stylaria lacustris. FIGURE 26 Perianal gills of Dero. Dero (Aulophorus) with long palps (right) and Dero
(Dero) without palps (left).
13b.
Dorsal chaetae either begin in III, hairs of III elongate “or” begin in II, the hairs being longest in midbody, and often missing from a
number of segments, and exceptionally thin when present...................................................................................................Bratislavia
14a(11b).
Posterior end of the body with a branchial fossa, normally with gills (Fig. 26); some species symbionts in tree frogs, these develop
gills once outside their host. ........................................................................................................................................................Dero
[There are three subgenera, Allodero containing the symbiotic species, Aulophorus with long palps on each side of the branchial
fossa, and Dero that lacks palps. In a strict cladistic sense, Allodero would not be recognized.]
14b.
No gills; free-living species ...............................................................................................................................................................15
15a(14b).
Dorsal chaetae begin in XVIII – XX, each bundle with a single short hair and a robust needle .............................................................
................................................................................................................................................Haemonais waldvogeli Bretscher, 1900
15b.
Dorsal chaetae begin in V, VI, or VII (note that ventral chaetae of IV and / or V may be missing).....................................................16
16a(15b).
Hair chaetae up to 9 per bundle, thick, with long lateral hairs (Fig. 27)..........................................................................Vejdovskyella
16b.
Hair chaetae 1 – 3 per bundle at most, thin with or without fine lateral hairs ...................................................................................17
17a(16b).
One to three especially long hair chaetae in each dorsal bundle of VI, 1 or 2 ordinary hair chaetae in the remainder (Fig. 28) ............
.............................................................................................................................................Slavina appendiculata (d’Udekem, 1855)
17b.
No elongate hair chaetae ..................................................................................................................................................................18
18a(17b).
Hair chaetae very short (84 – 120 m), present in only one or two bundles, some with no hair chaetae at all (see 6) ...........Piguetiella
18b.
Hair chaetae mostly 1 or 2 per bundle (two species with up to 5), long (180 – 200 m or more), present in all bundles from V or VI .
.........................................................................................................................................................................................................19
19a(18b).
Characteristic needle chaetae with thick, unequal teeth, sometimes clearly pectinate (Fig. 29); ventral chaetae change form and
length slightly between V and VI .............................................................................................................................................Allonais
452
Ralph O. Brinkhurst, and Stuart R. Gelder
FIGURE 27 Hair chaetae of Vejdovskyella, detail to right. FIGURE 28 Dorsal bundles of VI in Slavina
with elongate hairs. FIGURE 29 Needle chaetae of various Allonais species.
19b.
Needle chaetae simple-pointed, bifid, or faintly pectinate; either ventral chaetae of II differ from the rest “or” those of II – V differ
markedly in length, width, and form from the rest ...........................................................................................................................20
20a(19b).
Needle chaetae bifid or faintly pectinate; ventral chaetae of II may differ from the rest ..........................................................Specaria
20b.
Needle chaetae simple-pointed, bifid, or pectinate; ventral chaetae of II – V differ strongly in length, thickness, and form from the
rest in the majority of species. .......................................................................................................................................................Nais
[Species of Dero with the posterior end of the body missing may key out here because there will be no gills. Compare specimens to
those with gills; the anterior ventral chaetae in Dero species have very long upper teeth.]
D. Taxonomic Key to Genera and Selected Species of
Freshwater Tubificidae
This key is designed to maximize the use of external
characters. As it is primarily a key to genera, it is necessary to refer mostly to characters found on mature specimens. Immature specimens can often be identified on the
basis of chaetal form alone, but then the key must proceed directly to the specific level (Kathman and
Brinkhurst, 1998). In this key, there are two genus groups
that are difficult to separate on external characters; and
where this is true, evidence from distribution patterns is
introduced as an additional aid. Keys for worms of this
family are traditionally written at the species level.
1a.
Single dorsal and ventral gill filaments on posterior segments (Fig. 30) ........................................Branchiura sowerbyi Beddard, 1892
[Broken anterior fragments may be mistaken for Aulodrilus pluriseta (see 18b).]
1b.
No gill filaments .................................................................................................................................................................................2
2a(1b).
Body wall papillate (with projections usually covered with foreign matter attached by secretions) (Fig. 31).........................................
...............................................................................................................................Telmatodrilus (in part), Spirosperma, Quistadrilus
[Telmatodrilus (Alexandrovia) onegensis Hrabe, 1962 is recorded from Alaska; the subgenus is often elevated to generic status,
which may prove correct.]
2b.
Body wall naked.................................................................................................................................................................................3
3a(2b).
Spermathecal chaetae replace normal ventral chaetae in the spermathecal segment, very rarely in adjacent segments also (Fig. 34);
with or without modified penial chaetae (Figs. 39, 41, 42) and cuticular penis sheaths (Fig. 35) ........................................................4
3b.
No modified spermathecal chaetae .....................................................................................................................................................8
4a(3a).
Thin, hollow-tipped spermathecal chaetae in X, similar chaetae in XI together with apparent cuticular penis sheaths (Fig. 32)...........
............................................................................................................................................................Haber speciosus (Hrabe, 1931)
12. Annelida: Oligochaeta, Including Branchiobdellidae
453
4b.
Spermathecal chaetae not duplicated (or, if so, on VII or VIII and very rarely scattered from VI – XII); with or without cuticular
penis sheaths) .....................................................................................................................................................................................5
5a(4b).
Spermathecal chaetae broad, spatulate, one each side of IX associated with long tubular glands; ventral chaetae of X similar or absent; several penial chaetae in each ventral bundle of XI with short, knobbed distal ends; male ducts open into large median inversion of the body wall of XI (Fig. 33) ...................................................................................................Rhizodrilus lacteus Smith, 1900
5b.
Without this unique set of characters combined .................................................................................................................................6
6a(5b).
Spermathecal chaetae much broader and longer than the normal ventral chaetae, distal ends hollow, located on X (or in one
instance on VII or VIII or even irregularly between VI and XII); no cuticular penis sheaths .............................................Potamothrix
FIGURE 30 Branchiura, showing posterior fan of gill filaments. FIGURE 31 Papillate body wall of Spirosperma. FIGURE 32 Spermathecal chaeta of Haber (in which penial chaetae similar.) FIGURE 33 Reproductive organs, left side (anterior to left) of Rhizodrilus lacteus. Note spermathecal chaeta and gland (g) in IX, spermatheca (sp) in X, atrium (a) in XI discharging into median chamber that also receives penial chaetae (on right,
opposite atrial pore). FIGURE 34 Spermathecal chaeta of Isochaetides. FIGURE 35 Penis sheath of Isochaetides. FIGURE 36 Inverted (left) and everted penis of Psammoryctides. FIGURE 37 Varichaetadrilus, penis
(left) with penial sheath on tip (shown pointing down) and penial chaetae, top right and (enlarged) to right.
454
Ralph O. Brinkhurst, and Stuart R. Gelder
[Most species are restricted to the Great Lakes basin or extend into adjacent states; only one is widespread. These may be introduced from Europe, where they are characteristic of productive waters.]
6b.
Spermathecal chaetae longer than the normal ventral chaetae, slender distally, somewhat like a hypodermic needle (Fig. 34); true or
apparent cuticular penis sheaths present.............................................................................................................................................7
7a(6b).
True cuticular penis sheaths present, but not much thicker than the normal cuticular layer of the body wall (Fig. 35)......Isochaetides
7b.
Apparent penis sheaths present (probably the cuticular linings of eversible penes), look like crumpled cylinders in whole-mounts of
most specimens (Fig. 36) .............................................................................................................................................Psammoryctides
[One species is a recent introduction to the St. Lawrence River (Quebec) from Europe, another is known only from coastal swamps
of the Gulf of Mexico, and a third is known from California and the Great Lakes.]
8a(3b).
Penial chaetae present replacing the normal ventral chaetae of XI (Figs. 37, 39, 41, 42) ....................................................................9
8b.
Penial chaetae absent, ventral chaetae of XI usually missing in mature specimens ............................................................................12
9a(8a).
Penial chaetae bifid but with shortened distal ends and elongate proximal ends, somewhat thicker than the normal ventral chaetae;
with short penis sheaths on the ends of erectile penes (Fig. 37) .....................................................................Varichaetadrilus (in part)
[Distributed form Alaska to California and Alberta (but see also 14).]
9b.
Penial chaetae strongly modified, with knobbed or hooked tops to short distal ends, elongate proximal ends, arranged fanwise with
heads close together (as if they function as claspers) “or” large, single and sickle-shaped (Figs. 39, 41, 42) .....................................10
10a(9b).
Male pores open into an eversible chamber in XI that contains so-called paratria (Fig. 38) and penial chaetae when present (Fig.
39); sensory pit on the prostomium (Fig. 3); sperm attached to the body wall close to the male pore in external spermatophores
(Fig. 40)............................................................................................................................Bothrioneurum vejdovskyanum Stolc, 1888
10b.
No median copulatory chamber or sensory pit; sperm loose in spermathecae after copulation.........................................................11
11a(10b).
Penial chaetae in XI, 3 – 6 simple-pointed chaetae with hooked tips (Fig. 41); somatic chaetae 3 – 5 bifids from II – XX, simplepointed chaetae from XXV – XXX posteriad, no dorsal hair chaetae ............................Phallodrilus hallae Cook and Hiltunen, 1975
11b.
Penial chaetae with knobbed tips, or single, sickle-shaped (Fig. 42); no simple-pointed somatic chaetae (except in the ventral
bundles of one species, when accompanied by bifid chaetae, and the dorsal bundles of that species possess hair chaetae)....................
Rhyacodrilus
12a(8b).
Cuticular penis sheaths present in XI (Figs. 42 – 45) .........................................................................................................................13
12b.
Cuticular penis sheaths absent..........................................................................................................................................................15
13a(12a).
Penis sheaths more or less elongate, cylindrical, with penes free within them (Fig. 43); all chaetae bifid ...........................Limnodrilus
13b.
Penis sheaths annular to conical, attached to the surface of the penis (Figs. 44 and 45) ...................................................................14
14a(13b).
Short cuticular penis sheaths, annular to tub-shaped (Fig. 44)..........................Varichaetadrilus (in part), Tubifex, Ilyodrilus (in part)
[The Varichaetadrilus species included here are reported from the southeastern United States; I. frantzi Brinkhurst, 1965 is restricted to freshwater at the head of estuaries from British Columbia (Canada) to California.]
14b.
Penis sheaths conical (Fig. 45)...........................................................................Tasserkidrilus, Ilyodrilus templetoni (Southern, 1904)
15a(12b).
Chaetae simple-pointed anteriorly, behind the clitellum the chaetae have brushlike tips (Fig. 46) ...................................Telmatodrilus
vejdovskyi Eisen, 1879 [This species is known from British Columbia (Canada) to California.]
15b.
Chaetae bifid, or with upper tooth divided, or pectinate to palmate, all with the teeth in a single plane...........................................16
16a(15b).
Pharynx and mouth enlarged, prostomium reduced (Fig. 47); chaetae of III broad, curved, with large lower teeth, the rest thinner,
straighter, and with teeth more nearly equal; no spermathecae, presumably parthenogenetic ..Teneridrilus mastix (Brinkhurst, 1978)
16b.
Pharynx and mouth not enlarged, prostomium well developed; chaetae of III not modified; with spermathecae ..............................17
17a(16b).
Ventral chaetal bundles with up to 4 simple-pointed or bifid chaetae with reduced upper teeth. [In this species from Lake Tahoe
there are apparently no penial chaetae, although these are usually present in this genus (see 11b).] .....................................................
......................................................................................................................................Rhyacodrilus brevidentatus Brinkhurst, 1965
17b.
No simple-pointed ventral chaetae ...................................................................................................................................................18
18a(17b).
Dorsal chaetal bundles with 2 – 4 pectinate-to-palmate chaetae from II – XIV, from there posteriad all dorsal chaetae bifid; anterior
ventral chaetae 3 – 5 per bundle, bifid with the upper teeth thinner than but only a little longer than the lower, but beyond XIV the
upper teeth become much longer than the lower..............................................Arctodrilus wulikensis Brinkhurst and Kathman, 1983
[This species is recorded from the Brooks Range, Alaska.]
18b.
Chaetae progressively enlarged from III posteriad, becoming very broad with large lower teeth and recurved distal ends, “or” no
hair chaetae dorsally and large numbers of bifid chaetae with thin, short upper teeth, becoming palmate beyond VI in one species,
“or” hair chaetae present but all ventral chaetae again characteristically numerous and with short upper teeth (Fig. 48 .....Aulodrilus
12. Annelida: Oligochaeta, Including Branchiobdellidae
FIGURE 38 Paratrium of Bothrioneurum, shown left in situ in everted terminal portion of male duct, detail
to right. FIGURE 39 Penial chaetae, Bothrioneurum, detail of tip above, bundle below. FIGURE 40 Spermatophores of Bothrioneurum, attached to body wall around gonopores (left) and detail (right). Note
sperm enclosed, with no duct through attachment stalk. FIGURE 41 Penial chaeta, Phallodrilus hallae.
FIGURE 42 Penial chaetae, Rhyacodrilus falciformis. FIGURE 43 Penes of various Limnodrilus species,
three to right much longer than shown (40 times the length when fully developed). FIGURE 44 Penis sheath
of Tubifex tubifex, often thin and difficult to see, but granular surface often a clue. FIGURE 45 Penis sheath
of Ilyodrilus, short conical form. The terminal opening often looks torn, and individuals with a second portion
as long as that illustrated here may be seen, this second part presumably shed before mating. FIGURE 46
Posterior brushlike chaetae, Telmatodrilus. FIGURE 47 Inverted (above) and everted pharynx of Teneridrilus
mastix. FIGURE 48 Ventral chaeta, Aulodrilus.
455
456
Ralph O. Brinkhurst, and Stuart R. Gelder
BRANCHIOBDELLIDAE
VI. INTRODUCTION TO BRANCHIOBDELLIDAE
Branchiobdellids are leechlike, ectosymbionts living primarily on astacoidean crayfish and a few other
crustaceans. A single host can be infested with over
1,000 individuals and may include a taxonomic diversity of up to seven species representing five genera.
Branchiobdellids have an Holarctic distribution with a
total of 21 genera and 149 species (Gelder, 1996a), of
which 15 genera containing 107 species are reported
from North America. In spite of being known for almost 250 years, the ecophysiological interactions of
this complex ectosymbiotic association remain largely
unstudied.
Branchiobdellids have for the most part been considered to be a family, or at best a superfamily, until
Holt (1965) raised them to an order, Branchiobdellida.
He did this to make the taxon independent of, and
equal in rank to, both the oligochaetes and leeches. In
a review of the various ranks and names given to the
branchiobdellids (Gelder, 1996a), it was found that
the taxon has reached class rank, Branchiobdellae, but
this was an overly inflated rank for the group. Holt
(1986) arranged the branchiobdellid genera into five
families within the order. This gave rise to the term,
used in the first edition, “branchiobdellid” as the com-
mon epithet for members of the order, thus leaving
branchiobdellid for members of that family. The demotion of the taxon from order to family, and the five
families created by Holt to subfamily rank, enables the
relative relationships of these taxa to be maintained.
There is no single publication dealing with all of the
known species of branchiobdellidans, but a detailed
bibliography of the taxon was made by Sawyer
(1986). Therefore, as this work will deal only with
North American genera, citations are included
through which species descriptions and further information can be found.
VII. ANATOMY AND PHYSIOLOGY
OF BRANCHIOBDELLIDS
A. External Morphology
Adult branchiobdellids range from 0.8 – 10 mm in
length. The body in most species is rod-shaped (terete),
although some have a characteristic pyriform or flaskshape with either ventral or dorso-ventral flattening.
The segment number is constant and, based on the
number of paired ganglia, is accepted to be fifteen (XV)
as shown by Roman numerals in Figure 49. The external morphology shows a peristomium and three segments forming a head, and the remaining obvious body
FIGURE 49 Diagram of the lateral aspect of a hypothetical branchiobdellidan showing anatomic characters
used in the taxonomy of this group: a, anus; aa, anterior attachment surface; b, bursa; d, digitiform dorsal appendages; ds, developing sperm; e, esophagus; ga, glandular atrium; i, intestine; j, jaw; l, lip; m, mouth; ma,
muscular atrium; n, nephridia; np, nephridial pore; o, ovum; op, oral papillae; pa, posterior attachment surface; pg, prostate gland; ph, pharynx; s, stomach; sl, supernumerary longitudinal muscles; sp, spermatheca; su,
sulcus; t, tentacle; vd, vasa deferentia; H, head; P, peristomium; 1-11, body segments; I-XV, segments based on
paired ganglia.
12. Annelida: Oligochaeta, Including Branchiobdellidae
segments are numbered in Arabic numerals, 1 to 11.
The latter nomenclature is the one used in the descriptive literature on branchiobdellids. Segment 11 is modified into the posterior, disk-shaped attachment organ
(sucker), and contains a single pair of ganglia, XV.
Peristomial tentacles or lobes may be present on the
dorsal lip, and the mouth is surrounded usually by 16
oral papillae. Each body segment is divided into a major anterior and a minor posterior annulus, although a
few species have the major annulus further subdivided
into two. Dorsal transverse ridges across the major annuli are produced by supernumerary longitudinal muscles (sl) and sometimes support dorsal digitiform or
multibranched appendages (d) (Gelder, 1996b).
B. Organ System Function
The alimentary canal consists of a pair of sclerotized jaws (j) situated in the anterior pharynx followed
by one to three pairs of sulci (su), a short esophagus
(e), stomach (s), intestine (i), and an anus (a) opening
dorso-medially onto segment 10 (Fig. 49). Locomotory
movements are leechlike and involve an anterior and a
posterior attachment organ (Gelder and Rowe, 1988).
The anterior attachment site is on the ventral surface
of the ventral lip and is not the mouth as assumed generally. The muscle cells have the nucleus and cytoplasm in the medulla with the contractile fibers in the
cortex. The ultrastructure of the cells in three species
from the Branchiobdellinae are all round circomyarian
in their contractile fiber arrangement. A species from
each of the Bdellodrilinae and Cambarincolinae have a
few round circomyarian fibered cells, but most are
“polyplatymyarian” (Valvassori et al., 1994). The
pore(s) of the asymmetrically arranged anterior pair of
nephridia open to the exterior on segment 3 (np) and
the posterior pair are located in segment 8, but open
on segment 9.
A pair of testes are located in segments 5 and 6,
and the sperm develop (ds) in the coelom of those segments (Fig. 49). The ultrastructure of spermatozoa
from four species representing the Branchiobdellinae,
Bdellodrilinae, and Cambarincolinae showed an unexpected variation in morphology (Ferraguti and Gelder,
1990). Additional studies have confirmed that morphological variations in the spermatozoa occur between
genera, but that interspecific variations are dimensional
only (Ferraguti, 2000). Two vasa efferentia, both with
a ciliated funnel, merge into a vas deferens and the two
vasa deferentia, one from each segment, enter the glandular atrium in segment 6. When present, the prostate
gland forms a protuberance or a diverticulum from the
glandular atrium. The connecting muscular atrium
passes into the penis which is surrounded by a muscu-
457
lar bursa (b). Most penes are either eversible or protrusible, although a few species have a penis that shows
varying degrees of reduction. The bursal atrium opens
mesad on the ventral surface of segment 6. The terms,
“spermiducal gland” ( glandular atrium) and “ejaculatory duct” ( muscular atrium) as used in the First
Edition (Brinkhurst and Gelder, 1991) followed Holt
(1986). These have been replaced by the terms in
parentheses to promote consistency in anatomical terminology with oligochaetologists in general.
The spermatheca (sp), when present, is located in
segment 5 and consists of a glandular ental region connected to a muscular duct that opens medially onto the
ventral surface of the segment. A pair of ovaries in segment 7 produce eggs that mature in the coelom, and
are released through a pair of short ducts in the body
wall of that segment.
The vascular system consists of a dorsal and ventral longitudinal vessels that are connected by four
paired, lateral branches in the head and a single pair of
branches in segments 1, 7, and 10, respectively. The
dorsal vessel becomes a sinus surrounding the gut epithelium from segments 3 – 7.
The central nervous system consists of a dorsal
brain, circumesophageal connectives and a paired ventral cord extending into the last segment. The head
contains four pairs of ganglia with each body segment
containing a single pair of ganglia. The nerve cord is
displaced laterally in segments 5 and 6 where the spermatheca and male genitalia pass mesad through the
ventral body wall.
C. Environmental Physiology
Most branchiobdellids browse on the epibionts,
such as diatoms and ciliates, attached to the host and
are opportunistic commensals when fragments of the
host’s food are available (Jennings and Gelder, 1979).
Other opportunistic food sources include cannibalism
and exposed host tissues at injury sites. Contrary to
the reputation of these worms, a parasitic regime has
been demonstrated only in Branchiobdella hexodonta,
and inferred in a few other species. Food is sucked into
the mouth and then the jaws close to crop microorganisms from the substrate or fragment large particles.
Various mucoid secretions assist the ingestion of food,
but the term “salivary gland” as quoted by Sawyer
(1986) is inappropriate and was corrected by Gelder
and Rowe (1988). Digestion is extracellular in the
stomach and intestine with uptake occurring in the
cells of the latter region. Low oxygen tensions do not
appear to affect branchiobdellids adversely, although
some species tolerate higher water temperatures better
than others.
458
Ralph O. Brinkhurst, and Stuart R. Gelder
VIII. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
The branchiobdellids are characterized by a lack of
anatomical and ecological diversity in comparison with
equivalent clitellate taxa. The range of the branchiobdellids in North America extends from southern
Canada to Costa Rica (Gelder, 1999). A few species are
found distributed throughout this region, but the others have a more limited range (see Section IX.B). The
zoogeography of species is becoming more confused
and obscure as crayfish and their symbionts are introduced into new areas for aquaculture, live food, research, and teaching purposes.
B. Reproduction and Life History
Information on chromosome numbers is restricted
to three species, Branchiobdella astaci 2n 14, B.
kozarovi 2n 12, and B. parasita 2n 10 (Mihailova
and Subchev, 1981), and they refute previous reports of
n 4 and 8 for the first species. Germ cells from the
testes are released into the coelom of segments 5 and 6,
where spermatogenesis occurs. The mature spermatozoa then congregate at the mouth of the sperm funnels
prior to their passage to the penis. Prior to copulation,
the muscular bursa is partially or completely everted
(Figs. 50 – 53), and comes close to, or contacts with,
the epidermis surrounding the recipient’s spermathecal
pore. This is followed by the extended penis entering
the pore for insemination. Single inseminations have
been observed; however, it is not known if reciprocal
insemination occurs. Spermatophores on the epidermis
of Xironogiton victoriensis have been examined by
FIGURE 50
Gelder (1996c). Members of Ellisodrilus and
Caridinophila do not have spermatheca so their
method for sperm reception and storage is open to
speculation. Although the use of spermatophores has
been proposed as a solution to their problem, examinations of preserved specimens of Ellisodrilus have found
no such packets.
The secretions from the clitellar gland cells have
been characterized histochemically and compared with
those in other clitellate annelids (Gelder and Rowe,
1988). These secretions produce the pedunculate cocoon and nourishment to sustain the embryo while it
develops. Branchiobdellid cocoons deposited on live
hosts will develop to maturity and hatch out. The
hatching rate as observed on live hosts is temperature
dependent, and occurs after 10 – 12 days at 20 – 22°C.
The embryo is lecithotrophic and then becomes an albuminotrophic cryptolarva. In this latter phase, precocious feeding of the surrounding nutrient fluid starts
with ingestion by a transitory pharynx and the digestion yolk-sac epithelium. There are no reports of
growth rates or the time a juvenile requires to reach
sexual maturity. A few reports have noted that cocoons
were deposited on cast exoskeletons of their hosts and
the glass walls of their container, but embryos did not
develop and emerge. However, Woodhead (1945) is the
only person to report that branchiobdellids mated, attached their cocoons to the walls of the stock dishes,
and young hatched a few days later.
C. Ecological Interactions
Branchiobdellids have been reported on freshwater
crustaceans, primarily astacoidean crayfish. Although all
Diagrammatic section of a withdrawn, eversible penis: b, bursa, p, penis. FIGURE 51 Diagrammatic section of an extended, eversible penis; b, bursa; p, penis. FIGURE 52 Diagrammatic section of a
withdrawn, protrusible penis; b, bursa; p, penis; r, retractor muscle. FIGURE 53 Diagrammatic section of an
extended, protrusible penis: b, bursa, p, penis; r, retractor muscle.
12. Annelida: Oligochaeta, Including Branchiobdellidae
crayfish appear to be potential hosts, some species actively remove any branchiobdellids from their exoskeleton (Gelder and Smith, 1987). Noncrayfish hosts were
reported to be acceptable in troglodytic habitats (Holt,
1973b) and in Central America at and beyond the
southern limits of the geographical range of the astacoidean crayfish hosts (Gelder, 1999). Branchiobdellids
have been reported on the blue crab, Callinectes sp., in
low salinity waters adjacent to the Gulf of Mexico
(Overstreet, 1983), and so the possibility exists that a
marine branchiobdellid will be found.
Branchiobdellids also harbor ectosymbionts and
parasites. Externally, these include stalked, solitary and
colonial ciliate protozoans, and rotifers. Internally,
branchiobdellids serve as host to the third juvenile
stage of the nematode Dioctophyme renale (giant kidney worm) (Schmidt and Roberts, 1989), and the
trematode mesocercaria of Pharyngostomoides procyonis (reviewed in Schell, 1985).
D. Evolutionary Relationships
The branchiobdellids form a monophyly (Gelder
and Brinkhurst, 1990). The leechlike appearance of
branchiobdellids has caused some researchers to included them with the leeches. However, the presence of
a semiprosopore male ducts (Fig. 6) in both the branchiobdellids and lumbriculid oligochaetes, strongly
supports the branchiobdellids returning to family rank
as a sister group to the lumbriculids (Brinkhurst,
1999). The anatomical modifications shown by branchiobdellids, such as constant segment number, loss of
chaetae, development of anterior and posterior attachment organs, jaws, and other features, are not considered homologous to similar states in leeches but, instead, reflect a specialization to an ectosymbiotic
lifestyle as proposed by Stephenson (1930). The relocation of the branchiobdellids back into the oligochaetes
is the result of the best interpretation of the available
data. Phylogenetic analyses will soon be available using
matrices of nucleotide sequences in selected genes from
a significant number of clitellate species. Whether the
new methodologies will provide new phylogenies or
confirm some of those already established remains to
be seen.
459
IX. IDENTIFICATION OF BRANCHIOBDELLIDS
A. Collecting, Rearing, and Preparation
for Identification
Crayfish and other crustacean hosts are collected
by netting or baiting a trap (see Chapter 22) and then
placed directly into a container with preservative. If
branchiobdellids are to be examined it is recommended
that the preservative formalin – alcohol – acetic acid
(FAA) be used (70 mL ethanol 5 mL formaldehyde
(full strength) 1 mL glacial acetic acid 30 mL water). The volume of preservative should be at least three
times that of the crustaceans to ensure thorough preservation of the symbionts and host. Branchiobdellids removed from a live host have been maintained in vitro
for several months.
Microscopical examination of a living branchiobdellid— in water slightly squeezed between slide and
cover-glass — will enable all of the anatomical characters to be observed. First examine the specimen from a
dorsal or ventral aspect as this enables one to observe
the peristomium, jaws and location of the anterior
nephridial pore(s). The specimen can then be rolled to
show a lateral aspect, allowing the spermatheca and
male genitalia to be studied.
Live specimens can be relaxed using carbonated
drinks (such as club soda) or a 1 – 2% magnesium chloride solution (Delly, 1985), straightened or flattened,
and then immediately preserved. Unstained specimens
are dehydrated in graded ethanol solutions, cleared in
clove oil or oil of wintergreen (methyl salicylate), and
mounted in Canada balsam for permanent preparations. Specimens usually curl when preserved on the
host, and so these worms can only be examined from
the lateral aspect. Large branchiobdellids (4 – 10 mm
long) will require microdissection to expose their genitalia. The necessary instruments can be made from
fragments of a razor blade and insect (very small) pins
glued to the end of cocktail sticks. For techniques to
prepare serial sections and to stain specimens, refer to a
book on animal microtechiques, such as Humason
(1979).
Gelder (1996a) compiled a checklist of branchiobdellids and included citations to all papers containing
type descriptions.
B. Taxonomic Key to Genera of Freshwater Branchiobdellidae
1a.
Two anterior nephridial pores ............................................................................................................................................................2
1b.
One anterior nephridial pore ..............................................................................................................................................................4
2a(1a).
Ventral flattening of body...................................................................................................................................................................3
2b.
Dorso-ventral flattening of body (Fig. 54) .........................................................Xironogiton [British Columbia, Oregon, Washington,
California, Idaho, Wyoming, and the Appalachian Region from Maine to Georgia.]
460
Ralph O. Brinkhurst, and Stuart R. Gelder
FIGURE 54 Ventral view of Xironogiton instabilis, length about 2 mm. FIGURE 55 Lateral view of
Pterodrilus alcicornis, length about 1.5 mm. (Redrawn from Holt, 1986b.) FIGURE 56 Ventral view of
Triannulata magna, length about 4.5 mm. (Redrawn from Holt, 1974.) FIGURE 57 Lateral view of
Bdellodrilus illuminatus, length about 3 mm, lg, lateral glands. FIGURE 58 Lateral view of Ceratodrilus
thysanosomus, length about 3 mm. (Redrawn and modified from Holt, 1960.)
3a(2a).
Vasa deferentia entering glandular atrium mesad............................................................................Ankyrodrilus [Tennessee, Virginia]
3b.
Vasa deferentia entering glandular atrium entad................................................................................ Xironodrilus [East-central USA]
4a(1b).
Vasa deferentia entering glandular atrium mesad ...............................................................................................................................5
4b.
Vasa deferentia entering glandular atrium entad.................................................................................................................................7
5a(4a).
Body surface with indistinct segments ................................................................................................................................................6
5b.
Body surface with distinct segments (Fig. 59) ....................................................................................................Cronodrilus [Georgia]
6a(5a).
Lateral segmental glands, segments 1 – 9 (Fig. 55) ...............................................Bdellodrilus [Southern Canada to southern Mexico]
6b.
Lateral segmental glands not in segments 1 – 9 ...........................................................Uglukodrilus [California, Oregon, Washington]
7a(4b).
Spermatheca present...........................................................................................................................................................................8
7b.
Spermatheca absent ..........................................................................................Ellisodrilus [Tennessee, Kentucky, Indiana, Michigan]
8a(7a).
Penis protrusible (Figs. 52 and 53) .....................................................................................................................................................9
8b.
Penis eversible (Figs. 50 and 51) .......................................................................................................................................................11
9a(8a).
Prostate body present ......................................................................................................................................................................10
9b.
Prostate body absent (Fig. 60) ..................................................................................................................Magmatodrilus [California]
10a(8b).
Dorsal appendages present, range from paired digitiform unit to single transverse ridge on segment 8 (Fig. 56) .................................
Pterodrilus [Eastern United States]
10b.
Dorsal appendages absent (Fig. 61) ..........................................................................Cambarincola [Southern Canada to Costa Rica]
11a(9a).
Prostate body absent .......................................................................................................................................................................12
11b.
Prostate body present ......................................................................................................................................................................13
12a(11a).
Two annuli per segment (Fig. 62) ...........................................................................Sathodrilus (in part) [Florida, California, Mexico]
12b.
Three annuli per segment (Fig. 57) ................................................................................................Triannulata [Oregon, Washington]
13a(11b).
Dorsal appendages present (Fig. 58) ...........................................................Ceratodrilus [Utah, Wyoming, Idaho, Montana, Oregon]
13b.
Dorsal appendages absent. [This grouping cannot be resolved further without detailed histological preparation.](see Fig. 63) ............
Sathodrilus (in part) [South Carolina, Georgia, Mexico] (see Fig. 64) ..................Sathodrilus (in part) [Southern Canada to Mexico]
Oedipodrilus [Tennessee, Kentucky, Ohio, Pennsylvania, Indiana, Illinois].......................................................Tettodrilus [Tennessee]
12. Annelida: Oligochaeta, Including Branchiobdellidae
461
FIGURE 59
Lateral view of the male genitalia and spermatheca in Cronodrilus ogyguis. (Redrawn and
modified from Holt, 1968a.) FIGURE 60 Lateral view of the male genitalia and spermatheca in Magmatodrilus obscurus. (Redrawn and modified from Holt, 1967.) FIGURE 61 Lateral view of the male genitalia
and spermatheca in Cambarincola fallax. FIGURE 62 Lateral view of the male genitalia and spermatheca
in Sathodrilus hortoni. (Redrawn from Holt, 1973a.) FIGURE 63 Lateral view of the male genitalia and
spermatheca in Sathodrilus carolinensis. (Redrawn from Holt, 1968a.) FIGURE 64 Lateral view of the
male genitalia and spermatheca in Sathodrilus attenuatus. (Redrawn from Holt, 1981.)
LITERATURE CITED
Appleby, A. G., Brinkhurst, R. O. 1970. Defecation rate of three
tubificid oligochaetes found in the sediment of Toronto Harbour, Ontario. Journal of the Fisheries Research Board of
Canada 27:1971 – 1982.
Aston, R. J. 1984. The culture of Branchiura sowerbyi (Tubificidae,
Oligochaeta) using cellulose substrate. Aquaculture 40:89 – 94.
Bagheri, E. A., McLusky, D. S. 1984. The oxygen consumption of
Tubificoides benedeni (Udekem) in relation to temperature, and
its application to production biology. Journal of Experimental
Marine Biology 78:187 – 197.
Block, E. M., Moreno, G., Goodnight, C. J. 1982. Observations on
the life history of Limnodrilus hoffmeisteri (Annelida, Tubificidae) from the Little Calumet River in temperate North America.
International Journal of Invertebrate Reproduction 4:239 – 247.
Bonacina, C., Bonomi, G., Monti, C. 1987. Progress in cohort cultures of aquatic Oligochaeta. Hydrobiologia 155:163 – 169.
Bowker, D. W., Wareham, M. T., Learner, M. A. 1985. A choice
chamber experiment on the selection of algae as food and
substrata by Nais elinguis (Oligochaeta, Naididae). Freshwater
Biology 15:547 – 557.
Brinkhurst, R. O. 1965. Studies on the North American aquatic
Oligochaeta II. Tubificidae. Proceedings of the Academy of
Natural Sciences, Philadelphia 117:117 – 172.
Brinkhurst, R. O. 1974. The benthos of lakes. Macmillan, London.
Brinkhurst, R. O. 1982. Evolution in the Annelida. Canadian Journal
of Zoology 60:1043 – 1059.
Brinkhurst, R. O. 1989. A phylogenetic analysis of the Lumbriculidae (Annelida, Oligochaeta). Can. J. Zool. 67:2731 – 2739.
Brinkhurst, R. O. 1998. On the genus Eclipidrilus, including a
description of E. ithys n. sp. Canadian Journal of Zoology
76:644 – 659.
Brinkhurst, R. O. 1999. Lumbriculids, branchiobdellidans and
leeches, a review of progress. Hydrobiologia. (in press).
Brinkhurst, R. O., Austin, M. J. 1978. Assimilation by aquatic
oligochaetes. Internationale Revue der gesamten Hydrobiologie
63:863 – 868.
Brinkhurst, R. O., Chua, K. E. 1969. Preliminary investigation of the
exploitation of some potential nutritional resources by three
sympatric tubificid oligochaetes. Journal of the Fisheries Research Board of Canada 26:2659 – 2668.
Brinkhurst, R. O., Gelder, S. R. 1989. Did the lumbriculids provide
the ancestors of the branchiobdellids, acanthobdellids and
leeches? Hydrobiologia 180:7 – 15.
Brinkhurst, R. O., Jamieson, B. G. M. 1971. Aquatic Oligochaeta of
the world. Oliver and Boyd, Edinburgh, UK.
Brinkhurst, R. O., Chapman, P. M., Farrell, M. J. 1983. A comparative study of respiration rates of some aquatic oligochaetes in
relation to sublethal stress. Internationale Revue der gesamten
Hydrobiologie 68:683 – 699.
Chapman, P. M. 1981. Measurement of the short-term stability of interstitial salinities in subtidal estuarine sediments. Estuarine
Coastal and Shelf Science 12:67 – 81.
Chapman, P. M., Brinkhurst, R. O. 1981. Seasonal changes in interstitial salinities and seasonal movements of subtidal benthic
462
Ralph O. Brinkhurst, and Stuart R. Gelder
invertebrates in the Fraser River Estuary. Estuarine and Coastal
Shelf Science 12:49 – 66.
Chapman, P. M., Brinkhurst, R. O. 1984. Lethal and sublethal tolerances of aquatic oligochaetes with reference to their use as a
biotic index of pollution. Hydrobiologia 115:139 – 144.
Chapman, P. M., Brinkhurst, R. O. 1987. Hair today, gone tomorrow: induced chaetal changes in tubificid oligochaetes. Hydrobiologia 155:45 – 55.
Chapman, P. M., Farrell, M. A., Brinkhurst, R. O. 1982a. Effects
of species interactions on the survival and respiration of
Limnodrilus hoffmeisteri and Tubifex tubifex (Oligochaeta,
Tubificidae) exposed to various pollutants and environmental
factors. Water Research 16:1405 – 1408.
Chapman, P. M., Farrell, M. A., Brinkhurst, R. O. 1982b. Relative
tolerance of selected aquatic oligochaetes to individual pollutants and environmental factors. Aquatic Toxicology 2:47 – 67.
Chapman, P. M., Farrell, M. A., Brinkhurst, R. O. 1982c. Relative tolerances of selected aquatic oligochaetes to combinations of pollutants and environmental factors. Aquatic Toxicology 2:69 – 78.
Chatarpaul, L., Robinson, J. B., Kaushik, N. K. 1979. Role of tubificid worms on nitrogen transformations in stream sediment.
Journal of the Fisheries Research Board of Canada 36:673 – 678.
Christensen, B. 1984. Asexual propagation and reproductive strategies in aquatic oligochaetes. Hydrobiologia 115:91 – 95.
Clark, R. B. 1964. Dynamics in metazoan evolution. Clarendon
Press, Oxford.
Coates, K. A. 1989. Phylogeny and origins of the Enchytraeidae.
Hydrobiologia 180:17 – 33.
Delly, J. G. 1985. Narcosis and preservation of freshwater animals.
American Laboratory pp. 31 – 40.
Drewes, C. D., Fourtner, C. R. 1988. Hindsight and rapid escape in
an aquatic oligochaete. Page 278.5, in: Society for Neuroscience, Abstracts of the 18th Annual Meeting, Toronto, Ontario, November 13 – 18 1988. 14(1).
Ferraguti, M. 2000. Clitellata, in Adyiodi, K. G., Adyiodi, R. G., (Series Eds.), Reproductive biology of invertebrates. Jamieson, B.
G. M., (Vol. Ed.), Progress in male gamete ultrastructure and
phylogeny, Vol. IX, Pt B. Oxford & IBH Pub. Co., New Delhi,
India, pp. 125–182.
Ferraguti, M., Gelder, S. R. 1990. The comparative ultrastructure of
spermatozoa from five branchiobdellidans (Annelida: Clitellata). Canadian Journal of Zoology 69:1945 – 1956.
Fend, S. V, Brinkhurst, R. O. Hydrobiologia 428:1–59. A revision of
Rhynchelmis. (2000).
Gardner, W. S., Nalepa, T. F., Quigley, M. A., Malczyk, J. M. 1981.
Release of phosphorus by certain benthic invertebrates. Canadian Journal of Fisheries and Aquatic Science 38;978 – 981.
Gelder, S. R. 1996a. A review of the taxonomic nomenclature and a
checklist of the species of the Branchiobdellae (Annelida, Clitellata).
Proceedings of the Biological Society of Washington 109:653–663.
Gelder, S. R. 1996b. Description of a new branchiobdellidan species,
with observations on three other species, and a key to the genus
Pterodrilus (Annelida, Clitellata). Proceedings of the Biological
Society of Washington 109:256 – 263.
Gelder, S. R. 1996c. Histochemical characterization of secretions in
the reproductive systems of two species of branchiobdellidans
(Annelida, Clitellata): a new character for the phylogenetic matrix? Hydrobiologia 334:219 – 227.
Gelder, S. R. 1999. Zoogeography of branchiobdellidans (Annelida)
and temnocephalidans (Platyhelminthes) ectosymbiotic on
freshwater crustaceans, and their reactions to one another in
vitro. Hydrobiologia (406:21–31.).
Gelder, S. R., Brinkhurst, R. O. 1990. An assessment of the phylogeny of the Branchiobdellida (Annelida: Clitellata) using
PAUP. Canadian Journal of Zoology 68:1318 – 1326.
Gelder, S. R., Hall, L. A. 1990. Description of Xironogiton victoriensis n.sp. from British Columbia, Canada with some remarks
on other species and a Wagner analysis of Xironogiton (Clitellata: Branchiobdellida). Canadian Journal of Zoology
68:2352 – 2359.
Gelder, S. R., Rowe, J. P. 1988. Light microscopical and cytochemical
study on the adhesive and epidermal gland cell secretions of the
branchiobdellid Cambarincola fallax (Annelida: Clitellata).
Canadian Journal of Zoology 66:2057 – 2064.
Gelder, S. R., Smith, R. C. 1987. Distribution of branchiobdellids
(Annelida: Clitellata) in northen Maine, U.S.A. Transactions of
the American Microscopical Society 106:85 – 88.
Harman, W. J., Loden, M. S. 1978. A re-evaluation of the Opitocystidae (Oligochaeta) with descriptions of two new genera. Proceedings of the Biological Society of Washington 91:453 – 462.
Harper, R. M., Fray, J. C., Learner, M. A. 1981. A bacteriological investigation to elucidate the feeding biology of Nais variabilis
(Oligochaeta, Naididae). Freshwater Biology 11:227 – 236.
Holmquist, C. 1976. Lumbriculids (Oligochaeta) of Northern Alaska
and Northwestern Canada. Zoologische Jahrbucher (Systematik) 103:377 – 431.
Holt, P. C. 1960. The genus Ceratodrilus hall (Branchiobdellidae,
Oligochaeta) with the description of a new species. Virginia
Journal of Science 11:53 – 77.
Holt, P. C. 1965. The systematic position of the Branchiobdellidae
(Annelida: Clitellata). Systematic Zoology, 14:25 – 32.
Holt, P. C. 1967. Status of the general Branchiobdella and Stephanodrilus in North America with description of a new genus
(Clitellata: Branchiobdellida). Proceedings of the United States
National Museum 124:1 – 10.
Holt, P. C. 1968. New genera and species of branchiobdellid worms
(Annelida: Clitellata). Proceedings of the Biological Society of
Washington 81:291 – 318.
Holt, P. C. 1973a. A summary of the branchiobdellid (Annelida:
Clitellata) fauna of Mesoamerica. Smithsonian Contributions to
Zoology, No. 142:1 – 40.
Holt, P. C. 1973b. Branchiobdellids (Annelida: Clitellata) from some
eastern North American caves, with descriptions of new species
of the genus Cambarincola. International Journal of Speleology
5:219 – 256.
Holt, P. C. 1973c. Epigean branchiobdellids (Annelida: Clitellata)
from Florida. Proceedings of the Biological Society of Washington 86:79 – 104.
Holt, P. C. 1974. An emendation of the genus Triannulata Goodnight
1940, with the assignment of Triannulata montana to Cambrincola Ellis 1912 (Clitellata: Branchiobdellida). Proceedings of the
Biological Society of Washington 87:57 – 72.
Holt, P. C. 1981. New species of Sathodrilus Holt 1968, (Clitellata:
Branchiobdellida) from the Pacific drainage of the United States,
with the synonymy of Sathodrilus virgiliae Holt 1977. Proceedings of the Biological Society of Washington 94:848 – 862.
Holt, P. C. 1986. Newly established families of the order Branchiobdellida (Annelida: Clitellata) with a synopsis of the genera. Proceedings of the Biological Society of Washington 99:676 – 702.
Humason, G. L. 1979. Animal Tissue Techniques. 4th edition Freeman, San Francisco, CA.
Jamieson, B. G. M. 1981. The ultrastructure of the Oligochaeta. Academic Press, London.
Jennings, J. B., Gelder, S. R. 1979. Gut structure, feeding and digestion in the branchiobdellid oligochaete Cambarincola
macrodonta Ellis 1912, an ectosymbiote of the freshwater crayfish Procambarus clarkii. Biological Bulletin 156:300 – 314.
Karlckhoff, S. W., Morris, K. R. 1985. Impact of tubificid
oligochaetes on pollution transport in bottom sediments. Environmental Science and Technology 19:51 – 56.
12. Annelida: Oligochaeta, Including Branchiobdellidae
Kaster, J. L., Bushnell, J. H. 1981. Cyst formation by Tubifex tubifex
(Tubificidae). Transactions of the American Microscopical Society 100:34 – 41.
Kaster, J. L., Val Klump, J., Meyer, J., Krezoski, J., Smith, M. E.
1984. Comparison of defecation rates of Limnodrilus hoffmeisteri Claparéde (Tubificidae) using two different methods. Hydrobiologia 111:181 – 184.
Kathman, R. D., Brinkhurst, R. O. 1998. Guide to the freshwater
oligochaetes of North America. Aquatic Resources Center, College Grove, TN. 264 pp.
Krezoski, J. R., Robins, J. A., White, D. S. 1984. Dual radiotracer
measurement of zoobenthos-mediated solute and particle transport in freshwater sediments. Journal of Geophysical Research
89:7937 – 7947.
Lang, C. 1984. Eutrophication of Lakes Leman and Neuchatel
(Switzerland) indicated by oligochaete communities. Hydrobiologia 114:131 – 138.
Lazim, M. N., Learner, M. A. 1986a. The life-cycle and production
of Limnodrilus hoffmeisteri and L. udekemianus (Oligochaeta,
Tubificidae) in the organically enriched Moat-Feeder stream,
Cardiff, South Wales. Archiv fur Hydrobiologie (Supplement)
4:200 – 225.
Lazim, M. N., Learner, M. A. 1986b. The life-cycle and productivity
of Tubifex tubifex (Oligochaeta, Tubificidae) in the Moat-Feeder
stream, Cardiff, South Wales. Holarctic Ecology 9:185 – 192.
Leitz, D. M. 1987. Potential for aquatic oligochaetes as live food in
commercial aquaculture. Hydrobiologia: 155:309 – 310.
Lochhead, G., Learner, M. A. 1984. The cocoon and hatchling of
Nais variabilis (Naididae, Oligochaeta). Freshwater Biology
14:189 – 193.
Loden, M. S. 1981. Reproductive ecology of Naididae. Hydrobiologia 83:115 – 123.
Marian, M. P., Pandian, T. J. 1984. Culture and harvesting techniques for Tubifex tubifex. Aquaculture 42:303 – 315.
Mihailova, P. V., Subchev, M. A. 1981. On the karyotype of three
species of the family Branchiobdellidae (Annelida: Oligochaeta).
Comptes rendus l’Academie bulgare des Sciences 34:265 – 267.
Milbrink, G. 1983. An improved environmental index based on the
relative abundance of oligochaete species. Hydrobiologia
102:89 – 97.
Milbrink, G. 1993. Evidence for mutualistic interactions in freshwater oligochaete communities. Oikos 68: 317 – 322.
Moore, J. P. 1895. Pterodrilus, a remarkable discodrilid. Proceedings
of the Academy of Natural Sciences of Philadelphia pp.
449 – 454.
Nemec, A. F. L., Brinkhurst, R. O. 1987. A comparison of methodological approaches to the subfamilial classification of the Naididae (Oligochaeta). Canadian Journal of Zoology 65:691 – 707.
Newrlka, P., Mutayoba, S. 1987. Why and where do oligochaetes
hide their cocoons? Hydrobiologia 155:171 – 178.
Olsson, T. I. 1981. Overwintering of benthic macroinvertebrates in
ice and frozen sediment in a North Swedish River. Holarctic
Ecology 4:161 – 166.
Overstreet, R. M. 1983. Metazoan symbionts of crustaceans. in:
Provenzano A. J., Jr., Ed. The biology of Crustacea, pathobiology. Vol. 6, Academic Press, New York, pp. 155 – 250.
Poddubnaya, T. L. 1984. Parthenogenesis in Tubificidae. Hydrobiologia 115:97 – 99.
Sawyer, R. T. 1986. Leech biology and behaviour. Clarendon,
Oxford. pp. 1065.
463
Schell, S. C. 1985. Handbook of trematodes of North America,
North of Mexico. University of Idaho Press, Moscow, ID.
Schmidt, G. D., Roberts, L. S. 1989. Foundations of parasitology.
Times Mirror / Morsby, Boston, MA.
Stephenson, J. 1930. The Oligochaeta. Clarendon Press, Oxford.
Valvassori, R., de Eguileor, M., Lanzavecchia, G., Gelder, S. R. 1994.
Comparative body wall musculature and muscle fibre ultrastructure in branchiobdellidans (Annelida: Clitellata), and their
phylogenetic significance. Hydrobiologia 278:189 – 199.
Vincent, V., Vaillancourt, G., McMurray, S. 1978. Premiere mention
de Psammoryctides barbatus (Grube) (Annelida, Oligochaeta)
en Amerique du Nord et note sur sa distribution dans le haut
estuaire du Saint-Laurent. Le Naturaliste Canadien 105:77 – 80.
Wassell, J. T. 1984. Revision of the lumbriculid oligochaete
Eclipidrilus Eisen, 1881 with description of three subgenera and
Eclipidrilus (Leptodrilus) fontanus n. subgen. n. sp. from Pennsylvania. Proceedings of the Biological Society of Washington
97:78 – 85.
Wiederholm, T., Wiederholm, A. -M., Milbrink, G. 1987. Bulk sediment bioassays with five species of fresh – water oligochaetes.
Water, Air and Soil Pollution 36:131 – 154.
Woodhead, A. E. 1945. The life-history cycle of Dioctophyma renale,
the giant kidney worm of man and many other mammals. Journal of Parasitology, supplement 31:12.
Zoran, M. J., Drewes, C. D. 1987. Rapid escape reflexes in aquatic
oligochaetes: variations in design and function of evolutionary
conserved giant fiber systems. Journal of Comparative Physiology (A) 161:729 – 738.
ADDITIONAL SUGGESTED READINGS
In addition to the literature actually cited in the text, the following publications will provide more specific information on taxonomy
(Brinkhurst and Wetzel, 1984) and data derived from a series of international conferences on aquatic oligochaete biology.
Brinkhurst, R. O., Cook, D. G., Eds. 1980. Aquatic Oligochaete biology. Plenum Press, New York.
Brinkhurst, R. O., Wetzel, M. J. 1984. Aquatic Oligochaeta of the
World: Supplement. A Catalogue of New Freshwater Species,
Descriptions, and Revisions. Canadian Technical Report of Hydrography and Ocean Sciences 44:i-v 101 pp.
Bonomi, G., Erseus, C., Eds. 1984. Aquatic Oligochaeta. Developments in hydrobiology, No. 24. Junk, Dordrecht, Netherlands.
Brinkhurst, R. O., Diaz, R. J., Eds. 1987. Aquatic Oligochaeta.
Developments in hydrobiology, No. 40. Junk, Dordrecht,
Netherlands.
Coates, K. A., Reynoldson, T. B., Reynoldson, T. B., Eds. 1996.
Aquatic oligochaete biology. VI. Developments in hydrobiology,
No. 115. Junk, Dordrecht, Netherlands.
Kaster, J. L. (Ed.). 1989. Aquatic Oligochaeta. Developments in
Hydrobiology, No. 51. Junk, Dordrecht, Netherlands.
Healy, B., Reynoldson, T. B., Coates, K. A., Eds. 1999. Aquatic
oligochaete biology. VII. Developments in hydrobiology. Junk,
Dordrecht, Netherlands (in press.).
Reynoldson, T. B., Coates, K. A., Eds. 1994. Aquatic oligochaete
biology. VI. Developments in hydrobiology, No. 95. Junk,
Dordrecht, Netherlands.
13
ANNELIDA: EUHIRUDINEA
AND ACANTHOBDELLIDAE
Ronald W. Davies*
Fredric R. Govedich
Faculty of Science
Monash University
Clayton, Victoria 3168
Australia
Department of Biological Sciences
Monash University
Clayton, Victoria 3168
Australia
I. Introduction to Euhirudinea
and Acanthobdellidae
II. Taxonomic Status and Characteristics
III. Euhirudinea and Acanthobdellidae
A. General Anatomy
B. Physiology
IV. Ecology
A. Diversity
B. Foraging Relationships
C. Reproduction and
Life History
D. Dispersal
E. Predation
F. Behavior
G. Population Regulation
H. Ecotoxicology
I. INTRODUCTION TO EUHIRUDINOIDEA
AND ACANTHOBDELLIDA
Leeches form an important component of the benthos of most lakes and ponds, and some species commonly occur in the quieter flowing sections of streams
and rivers. There are a total of 73 species presently
recorded from North America, of which the majority
are predators feeding on chironomids, oligochaetes,
amphipods, and molluscs. The young of several species
feed on zooplankton, but Motobdella montezuma is
unique in specializing on planktonic amphipods using
mechanoreception to detect its prey. Other species of
leeches are temporary sanguivorous (blood feeding)
* Present address: Department of Biological Sciences,
University of Calgary, Calgary, Alberta, Canada T2N 1N4.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
V. Collection and Rearing
A. Collection
B. Rearing
VI. Identification
A. Preparation of Specimens
B. Important Features
for Identification
C. Family Glossiphoniidae
D. Family Piscicolidae
E. Family Hirudinidae
F. Family Erpobdellidae
VII. Taxonomic Keys
A. Euhirudinea and Acanthobdella
B. Species of Euhirudinea
Literature Cited
ectoparasites of fish, turtles, amphibians, crocodilians,
water birds, or occasionally humans. The majority of
predaceous leeches have an annual or biannual life cycle, breeding once and then dying (semelparity). However, at least two species have been shown to be genetically capable of breeding several times (iteroparity)
although exhibiting phenotypic semelparity in the field.
The majority of sanguivorous leeches are iteroparous
showing saltatory growth after the three or more blood
meals required to reach mature size.
Because of the diversity of physico-chemical conditions encountered in different aquatic ecosystems and
temporally within a single habitat, leeches show considerable physiological plasticity. They are able to live
in waters with very low salt concentrations as well as
waters with salinities that exceed the salinity of seawater. Similarly, some species are capable of surviving in
465
466
Ronald W. Davies and Fredric R. Govedich
the absence of oxygen (anoxia) for more than 60 days
and in the presence of supersaturated water (hyperoxia) for shorter periods.
In many small ponds and lakes, leeches are the
top predators and are ideal for studying intra-and
interspecific resource competition, niche overlap, and
predator – prey interactions. In larger lakes and rivers,
leeches may form an important component of the diet
of fish; hence, they are grown commercially for fish
bait. Sanguivorous leeches are being investigated extensively in relation to the pharmacological properties of
their salivary secretions, especially their anticoagulants.
The leech-like Acanthobdella peledina, a temporary ectoparasite of salmonids, is the only species of
Acanthobdellidae in North America. Its ecology and
feeding have not been extensively studied, but are likely
to be similar to the Piscicolidae — a family of leeches
also ectoparasitic on fish.
II. TAXONOMIC STATUS AND CHARACTERISTICS
There is an undisputed close taxonomic affinity between leeches and oligochaetes, although there are differences of opinion about the exact nature of the relationship. This has resulted in leeches being variously
classified as Hirudinea, Hirudinoidea, or Euhirudinea.
Purschke et al. (1993) showed that not only did the
Branchiobdellae (see Chapter 12), Acanthobdellidae,
and Euhirudinea resemble each other, but they were
also closely related. Their evolutionary relationship remains unanswered, but the present consensus is to consider them all superorders of the subclass Clitellata.
The Clitellata are thought to have evolved in the interstitial environment of lagoons near the tributary outlets
and entrance to channels opening into the sea in the
Palaeozoic, 500 million years ago. The first documented fossil of a leech dates back 150 million years to
the Jurassic. However, a large annulate with a circular
structure similar to a leech sucker discovered at
Waukesha, Wisconsin (Briggs, 1991), confirmed as a
leech, will extend their history back 400 million years
to the Silurian. Cocoons of extant leeches have been
recorded from Holocene sediments but Manum et al.
(1991) suggest that ovoid fossils 1.0 – 10.0 mm long
with a netted or a feltlike wall fabric from the Lower
Cretaceous (150 million years) were very probably
leech cocoons.
Only a few families of microdrilid oligochaetes can
be considered as possible close relatives of the leeches.
The naiids are the best candidates as most live in aerated freshwater, which seems to be the original environment of leeches, many of them are predators, they
have a reduced and sometimes fixed number of
FIGURE 1
Dorsal view of the general body shape of a member
of each of the families: (a) Glossiphoniidae; (b) Piscicolidae; (c)
Hirudinidae; and (e) Erpobdellidae.
segments, they have eyes of very similar structure to
those of leeches and at least one genus (Chaetogaster)
has epizoic species which move by looping and which
like leeches lack a prostomium (Omodeo, 1998).
The diagnostic features of leeches are: (1) 32 postoral metameres (Fernandez, 1980, Weisblat et al.,
1980) plus a nonsegmental prostomium; (2) a reduced
or obliterated coelom; (3) absence in the adult of
chaetae and septa; (4) the presence of an anterior (oral)
sucker and a usually larger posterior (anal) sucker
(Fig. 1); (5) superficial subdivision of the metameres
into annuli (Figs. 2 and 29a, b); and (6) a median, unpaired, male genital pore placed anterior to the single
median ventral, female genital pore (Figs. 3 and 4a).
The characteristics delimiting the Euhirudinea are
primarily related to their feeding and feeding behavior
indicative that leeches are oligochaetes which at first
become predators and then, subsequently, on at least
two separate occasions some became sanguivorous
(Siddall and Burreson, 1995).
The phylogenetic position of the Acanthobdellidae
has been debated. Because they share some morphological, behavioral, and ecological characteristics more
typically attributed to leeches some researchers have included them within the same taxonomic group as the
FIGURE 2
Annulation showing (a) leech 3-annulate condition and
(b) Acanthobdella peledina 4-annulate condition.
13. Annelida: Euhirudinea and Acanthobdellidae
FIGURE 3 Ventral view of the anterior male and posterior female
gonopores separated by (a) four annuli and (b) two annuli.
leeches (Mann, 1961, Elliott and Mann, 1979; Klemm,
1982, 1985; Sawyer, 1972, 1986). However, recent
research shows that the Acanthobdellidae, along with
the Branchiobdellidae, are sister groups to the leeches
sharing a common ancestor (Siddall and Burreson,
1995, 1996). Acanthobdellidae are temporary ectoparasites of salmonid fishes but, unlike leeches, with five
anterior coelomic compartments separated by distinct
septa, each anterior segments bearing two pairs of
chaetae (setae) (Fig. 4b). The Acanthobdellidae do not
FIGURE 4
Acanthobdella peledina, ventral view of (a) the male
and female gonopores with a spermatheca and (b) the anterior view
showing the absence of an anterior sucker, the presence of a mouth
pore, and five pairs of hooked chaetae.
467
have an anterior sucker, but a small posterior sucker is
formed from four of the 30 segments forming the body.
Each segment is superficially subdivided into four annuli (Fig. 2b).
Classically, oligochaetes and leeches are placed
within the phylum Annelida either in the order
Hirudinea, class Clitellata, or in the class Euhirudinea.
Soös (1965, 1966a, b, c, 1967, 1968, 1969a, b) placed
leeches in the taxonomically undefined Hirudinoidea,
but subsequently both Soös (1969b) and Richardson
(1969) referred them to the class Hirudinoidea. Correctly, Klemm (1985) and Sawyer (1986) indicated that
in accordance with the Zoological Code the suffix
“-oidea” should only be added to the stem name of a
superfamily and recommended the suppression of the
class Hirudinoidea. Davies (1971, and many subsequent references) classified leeches as Hirudinoidea implicitly as a superfamily of the class Oligochaeta.
Until recently, these differences were primarily academic because everyone agreed that as Hirudinea, Euhirudinea or Hirudinoidea, leeches were in the phylum
Annelida and closely related to oligochaetes. However,
Sawyer (1986) defined the class Hirudinea in a significantly different way, with Euhirudinea, Acanthobdellidae, Branchiobdellidae, and Agriodrilidae as subclasses
within the phylum Uniramia. Thus, the class Euhirudinea (sensu Sawyer, 1986) required the removal of
this class from the phylum Annelida, and its placement
into a phylum with the Onychophora, Myriapoda, and
Hexapoda. The evidence to support this is scant
(Sawyer, 1984, 1986) and contentious (Davies, 1987),
and the majority of authors did not follow this recommendation and retained the leeches as Euhirudinea,
Hirudinea or Hirudinoidea as a class of Annelida, a
view supported by Zrzavy et al. (1998). The term Euhirudinea used in this chapter refers to a class of Annelida containing the leeches and not the Uniramian class
defined by Sawyer (1986).
Sawyer (1986) combined many of the families and
genera described and accepted by previous researchers,
including those described in detail by Richardson
(1969, 1971, 1975). In addition, he placed the North
American members of the genus Dina into Erpobdella
leading to further confusion about the relationships
within the Erpobdellidae and, particularly, between the
genus Erpobdella and Mooreobdella. These changes
were not justified scientifically nor were complete descriptions of the modified families and genera given.
This has resulted in taxonomic confusion. Leech families and genera described prior to Sawyer (1986) have
been retained in this chapter and until a formal revision, including a re-analysis of the higher level relationships of leeches, has been published it is recommended
that these names be retained.
468
Ronald W. Davies and Fredric R. Govedich
III. EUHIRUDINEA AND ACANTHOBDELLIDAE
A. General Anatomy
The body of a leech consists of two preoral, nonmetameric segments and 32 postoral metameres designated I through XXXIV, while the Acanthobdellidae
have only 29 postoral metameres (designated I through
XXIX). Each segment is usually divided externally by
superficial furrows into 2 – 16 annuli (Figs. 2 and 29).
Segments with the full number of annuli (complete segments) are found in the middle of the body; because
this number is generally characteristic of the genus or
species, it is referred to in the keys. Annuli features can
be most easily seen in the lateral margins of the ventral
surface.
Moore (1898) numbered the three primary annuli
a1, a2, and a3 from the anterior, with annulus a2
(neural annulus) containing the nerve cord ganglion.
The neural annulus (a2) is marked externally by transverse rows of cutaneous sensillae (Figs. 2 and 29). Repeated bisection of the primary annuli gives more complex annulation.
The number and position of the eyes are also important diagnostic features. In some genera, coalescence of eyes sometimes occurs, but the lobed nature of
the eyes usually indicates the original condition (Fig.
5b). Eyespots (oculiform spots) can also occur on the
lateral margins of the body and on the posterior sucker
(Figs. 5c, 6b, and 19 b, c, d).
FIGURE 5 Dorsal view of the head of Batracobdella with a single
pair of eyes (a) close together; (b) lobed; and (c) Placobdella hollensis
with one pair of eyes followed by pairs of accessory eyes.
FIGURE 6 (a) Cystobranchus verrilli; and (b) Cystobranchus meyeri showing the pulsatile vesicles on the urosome and the presence on
C. meyeri of oculiform spots on the posterior sucker and the paired
lateral oculiform spots on the urosome.
Papillae, small protrusible sense organs, are often
widely scattered over the dorsal surface. Tubercles,
which are larger protrusions that include some of the
dermal tissues and muscles, may also be present and
can bear papillae.
The body of most leeches and acanthobdellids is
not divisible into regions. With the genera Illinobdella
and Piscicolaria as exceptions, the body of the family
Piscicolidae is divided into a narrow anterior trachelosome and a longer and wider posterior urosome (Figs.
1b and 6). Lateral gills are absent, but some genera of
Piscicolidae have paired pulsatile vesicles on the neural
annuli of the urosome (Figs. 1b and 6a, b).
The anterior sucker of leeches may be very prominent (Figs. 1b, 6, 7b) or simple (Figs. 1a, c, d, 7a, c),
consisting only of the expanded lips of the mouth. In
the acanthobdellids, anterior suckers are entirely absent
(Fig. 4b). The posterior sucker of a leech is generally
directed ventrally and is wider than the body at the
point of attachment to the substrate (Figs. 1, 6, 19, and
22 – 24). The mouth is either a small pore on the edge
(Fig. 7a) or center (Fig. 7b) of the ventral surface of the
anterior sucker, or is large and occupies the entire cavity (Fig. 7c) of the anterior sucker. In the family Hirudinidae, the buccal cavity (which may or may not
13. Annelida: Euhirudinea and Acanthobdellidae
469
FIGURE 9
Surface view of the jaws of Hirudinidae showing
the teeth arranged in: (a) one (monostichodont) or (b) two (distichodont) rows.
FIGURE 7
Ventral view of the anterior sucker of: (a) the mouth
pore on the anterior rim of the sucker (e.g., Marvinmeyeria, Placobdella, Oligobdella); (b) near the center of the sucker (Piscicolidae,
Batracobdella, Helobdella); and (c) the mouth occupying the entire
cavity (e.g., Hirudinidae, Erpobdellidae).
FIGURE 8
Ventral view of the dissection of the mouth and buccal
cavity of: (a) Mollibdella grandis; (b) Percymoorensis marmorata;
and (c) Macrobdella decora showing the velum, and the relative size
of the jaws in Macrobdella decora and P. marmorata and the absence
of jaws in Mollibdella grandis.
contain jaws) is separated from the cavity of the anterior sucker by a flap of skin (the velum) (Fig. 8). When
present, there are three muscular jaws (two ventrolateral and one dorso-medial) (Fig. 8b, c) bearing teeth
arranged in one (monostichodont) (Fig. 9a) or two (distichodont) rows (Fig. 9b).
In the Erpobdellidae, the buccal cavity contains
three muscular ridges, but in the Glossiphoniidae and
Piscicolidae, the pharynx is modified to form an eversible muscular proboscis. Opening into the pharynx are
the salivary glands which secrete some digestive enzymes. In bloodsucking leeches the salivary glands are
primarily related to the process of bloodsucking rather
than digestion. The components of the salivary secretions and their functions can be summarized as lubrication, proteolytic inhibition, spreading (hyaluronidase),
vasodilation, and anticoagulation (e.g., hirudin or hementin) (Sawyer, 1986). The anticoagulants produced
by North American sanguivores have not been investigated extensively with work so far concentrating on the
European Hirudo medicinalis and the South American
Haementaria ghilianii. Seymour et al. (1990) and
Krezel et al. (1994) reported the presence and structure
of decorsin, a potent inhibitor of platelet aggregation
from Macrobdella decora.
The pharynx in most erpobdellid leeches leads to a
simple tubelike crop that lacks lateral ceca. However,
this is not the case in the erpobdellid genus Motobdella
which has one or two pairs of simple ceca or postceca
located in the posterior portions of the crop (Davies
et al. 1985; Govedich et al. 1998). The crop in most
leech families is adapted for the storage of fluids and
usually has 1 – 11 pairs of lateral ceca. Between the last
pair of crop ceca, the intestine leads to the anus (Fig.
10a). Only in the Glossiphoniidae does the anterior
portion of the intestine give off four pairs of simple
ceca (Fig. 10b). The anus usually opens on the dorsal
surface on or near segment XXVII, just anterior to the
posterior sucker. In a few species, the anus is displaced
anteriorly. Motobdella has one or two pairs of post
ceca (Fig. 10c).
Digestion and absorption mainly occur in the intestine although the crop is also used in predatory species.
470
Ronald W. Davies and Fredric R. Govedich
FIGURE 10
Stylized ventral view of the alimentary canal of (a)
Hirudinidae; (b) Glossiphoniidae; (c) Motobdella; and (d) Acanthobdella peledina.
In sanguivores, digestion is primarily by enzymes
produced by endosymbiotic bacteria. The bacteria involved are species specific for each species of sanguivorous leech (Jennings and Vanderlande, 1967), e.g.
Hirudo medicinalis utilizes Aeromonas hydrophila
FIGURE 11
(Pseudomonas hirudinis) (Büsing, 1951, Büsing et al.,
1953, Jennings and Vanderlande, 1967). Endopeptidases are absent or rare in leeches, but exopeptidases
occur abundantly in predatory species and protease
(Zebe et al., 1986) and protease inhibitors have been
recorded (Roters and Zebe, 1992a, b). Because of their
dependence on endosymbiotic bacteria, digestion in
sanguivorous leeches takes weeks or months; but for
predatory species, digestion requires just a few days
(Davies et al., 1977, 1978, 1981, 1982a; Wrona et al.,
1979, 1981).
The alimentary canal of acanthobdellids (Fig. 10c)
is similar to that of the glossiphoniid leeches. A small
eversible proboscis leads into the muscular pharynx,
which, in turn, leads into a large tubular crop lacking
ceca. The crop leads into the intestine, which bears six
pairs of lateral ceca; the anus is located dorsally between segments XXV and XXVI. Digestion is probably
similar to sanguivorous leeches with the use of endosymbiotic bacteria.
The male and female gonopores are visible in
mature leeches on the midline of the ventral surface of
segments XI and XII, respectively, and are separated by
a species-specific number of annuli (Figs. 3, 4, and 27).
The anterior male gonopore is larger and more easily
seen than the female gonopore. Leeches do not have
true testes or ovaries, but instead possess testisacs and
ovisacs. The testisacs are located posterior to segment
XI and are either discrete, paired spherical structures
(Fig. 11a) (e.g., Hirudinidae, Glossiphoniidae, and
Reproductive systems of: (a) Hirudo medicinalis (Hirudinidae); and (b)
Nephelopsis obscura (Erpobdellidae).
13. Annelida: Euhirudinea and Acanthobdellidae
Piscicolidae) located intersegmentally or multifollicular
columns resembling bunches of grapes lying on either
side of the ventral nerve cord (Fig. 11b) (e.g., Erpobdellidae; Singhal and Davies, 1985; Davies et al., 1985,
Govedich et al., 1998). Short vasa efferentia connect
the testisacs to the vasa deferentia on each side of the
body, which run anteriorly to form large coiled epididymis (sperm vesicles). Paired ejaculatory ducts run
from the epididymis through the atrial cornua and
unite to form a medial atrium consisting of a bulb and
eversible penis. There is a single pair of ovisacs which
are either small, spherical organs confined to segment
XII or elongate, coiled tubes sometimes bent back on
themselves such that their blind ends lie close to the
female gonopore (Singhal and Davies, 1985; Davies
et al., 1985). Two oviducts run from the ovisacs to
converge and form the common oviduct leading to the
female gonopore (Fig. 11).
Leeches are frequently recorded as sequential
protandrous hermaphrodites (Van Damme, 1974;
Lasserre, 1975). Nephelopsis obscura shows sequential
protandry during its first bout of gametogenesis, although there is a short transition period when gametogenic stages from both sexes co-occur. In the second cycle of gametogenesis, mature sperm and ova occur
simultaneously (Davies and Singhal, 1988) and thus
this cycle demonstrates simultaneous hermaphroditism.
At the time of egg laying, a cocoon is secreted by the
clitellum, a specialized region of epidermis usually posterior to the gonopores. As the cocoon moves toward
471
and passes over the head, fertilized eggs pass into the
cocoon from the female gonopore.
The reproductive system of Acanthobdellida peledina is basically similar with a pair of elongate testisacs
and ovisacs and median unpaired male and female
gonopores.
The excretory system of leeches and acanthobdellids
consists of a maximum of 17 pairs of highly modified
metanephridia opening along the body between segments VII and XXII. Many species do not exhibit the
full number of metanephridia, showing reductions in the
more anterior metameres and / or the segments containing gonads. In addition to excreting nitrogenous wastes
(mainly ammonia), the nephridia help maintain water
and salt balance in the body. With the osmotic pressure
of the body higher than the surrounding freshwater,
there is a tendency for an inward flow of water through
the epidermis. The nephridia pass out excess water and
help retain inorganic ions. Only about 6% of the salts
found in the primary urine are excreted, the rest being
reabsorbed (Zerbst-Boroffka, 1975). During acclimation
to hypersaline media, several different active and passive
processes are responsible for the changes of osmolarity
and electrolytic concentrations in the blood of the leech
(Nieczaj and Zerbst-Boroffka, 1993).
In the leeches and acanthobdellids, the large
coelom, which in Annelida is typically divided by intersegmental septa, has to varying degrees been obliterated (Fig. 12). In the Glossiphoniidae (Fig. 12a),
Piscicolidae (Fig. 12d), and acanthobdellids, the blood
FIGURE 12 Stylized transverse sections of: (a) Glossiphoniidae; (b) Hirudinidae; (c) Erpobdellidae; and (d)
Piscicolidae showing the coelomic sinuses, dorsal and ventral blood vessels, and the double ventral nerve cord.
472
Ronald W. Davies and Fredric R. Govedich
vascular system consists of: (1) a ventral vessel enclosed
with the nerve cord in the ventral lacuna; (2) a dorsal
blood vessel enclosed in the dorsal lacuna; and (3)
transverse blood vessels connecting the dorsal and ventral vessels in the anterior and posterior parts of the
body. In the Hirudinidae (Fig. 12b) and Erpobdellidae
(Fig. 12c), the true blood vascular system is completely
lost; instead, the blood circulates in the coelomic
lacuna system, forming a hemocoel.
Oxygen uptake in freshwater leeches and acanthobdellids is through the general body surface, although some species of Piscicolidae have small pulsatile
vesicles filled with coelomic fluid that function as accessory respiratory organs. Most leeches can also ventilate
their body surfaces by dorsoventral undulations along
the body reminiscent of swimming, but with the posterior sucker firmly attached. However, Acanthobdella
peledina does not ventilate (Dahm, 1962).
All leeches and acanthobdellids show typical looping; locomotory movements consisting of body elongation and shortening, with the anterior sucker (leeches
only) and posterior sucker serving alternately as points
of attachment. Many leech species are also able to
swim using dorsoventral undulations. All Erpobdellidae and Hirudinidae are good swimmers, but only a
few species of Glossiphoniidae and Piscicolidae are efficient swimmers. Some species (Placobdella hollensis)
swim readily as juveniles and adults, but others (Placobdella ornata and Placobdella parasitica) swim only
as juveniles (Sawyer, 1981). The erpobdellid genus Motobdella is unique in its swimming ability and specializes in feeding almost exclusively on pelagic amphipods
by using mechanoreception to identify and track its
prey in the water column (Blinn and Davies, 1990;
Blinn et al., 1987, 1988, 1990; Davies et al., 1985).
B. Physiology
Perhaps the greatest variability in the abiotic environment experienced by leeches occurs in reference to
temperature and dissolved oxygen concentrations
In stratified lakes, the epilimnion is generally at or
above saturation during the summer, while in the hypolimnion oxygen depletion occurs. Under ice, especially at the substrate – water interface, hypoxia and
possibly anoxia occur (Babin and Prepas, 1985; Baird
et al., 1987a). Thus, hypoxia or anoxia can occur on a
diel basis during the open-water season and on a longterm basis during ice cover. As spring approaches, the
water temperature rises above 5°C, snow cover melts
and under-ice primary productivity can result in shortterm (1 – 2 days) hyperoxia. Similarly during the summer, long-term hyperoxia can persist for several days.
Both anoxia and hyperoxia can cause mortality of
vertebrates and invertebrates (Casselman and Harvey,
1975). In Europe, Mann (1956) showed that oxygen
uptake by some leech species was proportional to the
oxygen concentration of the water (i.e., they were
conformers), while other species (Glossiphonia complanata, Helobdella stagnalis) showed a comparatively
constant oxygen uptake over a range of dissolved oxygen concentrations of the water (i.e., they were regulators). Nephelopsis obscura and Erpobdella punctata
exposed to short-term hypoxia both showed a decline
in oxygen uptake with reduced oxygen tension in the
water (Wrona and Davies, 1984). When exposed to hypoxic conditions, most leeches, but not Acanthobdella
peledina, show ventilation movements with the posterior sucker attached to the substrate and the body
thrown into dorsoventral flexions (Dahm, 1962). It is
assumed that the primary function of ventilation is
respiratory since ventilation increases under conditions
of low oxygen availability. While N. obscura and E.
punctata ventilate in hypoxic conditions, ventilations
also occur under saturated (normoxic) conditions and
always cease when oxygen decreases below 20%
saturation. This suggests ventilation may have additional roles.
Many leeches can withstand anaerobic conditions
(Von Brand, 1944), but little research has been done on
the ecological effect of anoxia (or hypoxia) on the
leeches, with the majority of work related to use of
leeches as indicator species in the European saprobic
system (reviewed in Sladacek and Kosel, 1984).
In flow-through experiments, Davies et al. (1987)
showed inter- and intraspecific differences in the survivorship of N. obscura and E. punctata to anoxic conditions and demonstrated the errors of using static
experiments for this type of study. For both species,
survivorship decreased with increased water temperature. Large N. obscura had longer survival times ( 50
days) than small individuals (12 days) at 5°C and also
at 20°C (8 and 4 days, respectively). Conversely, at 5°C
large E. punctata had shorter survival times (25 days)
than small individuals ( 50 days), but at 20°C, large
individuals survived longer (8 days) than small individuals (2 days). As young (small) N. obscura have a low
probability of surviving anoxia at winter temperatures
(5°C), it is evident that large N. obscura would be taking less risk if they overwintered rather than hazarding
cocoon production and hatching of young up to or
close to ice formation. In contrast, large E. punctata,
which have a low probability of surviving winter
anoxia, would be taking less risk to continue cocoon
production later in the season since their offspring can
survive winter anoxic conditions. These results correlate well with the patterns of reproduction observed in
the field (Davies, 1978; Davies et al., 1977).
13. Annelida: Euhirudinea and Acanthobdellidae
The metabolic and physiological adaptations of N.
obscura to long-term anoxia have been studied under
laboratory conditions mimicking winter conditions
(Reddy and Davies, 1993a, b; Dratnal et al., 1993).
Lactic acid did not accumulate in either large or small
individuals over a 40-day period of anoxia; however,
succinate and alanine did. Both glycogen and amino
acids were utilized as energy reserves during the anoxic
period along with stored lipids. Following the 40-day
period of anoxia, glycolytic flux and adenosine triphosphate diphosphate (ATP / ADP) were significantly reduced in both small and large individuals. Upon a return to normoxia, preanoxic levels of glycogen, malate,
succinate, alanine, and aspartate are restored within 24
hs. In addition, the preanoxic energy charge is restored
within the first 6 hs of normoxia. All small and 98% of
large individuals survived the 40-day anoxic period
without significant losses in body weight; however, activity was reduced when compared to N. obscura not
exposed to anoxic conditions (Reddy and Davies,
1993a, b). Physiological adaptations to the effects of
severe hypoxia are similar to those observed for anoxic
conditions and in N. obscura include decreased
growth, enhanced energy allocation of glycogen,
triclyglycerols and total lipids. Body weight and size at
maturity were not affected by hypoxia; however, the
time to maturity was longer for N. obscura under hypoxic conditions (Dratnal et al., 1993).
Even less work has been done on the effect of hyperoxia on leeches. Survival of N. obscura and E.
punctata exposed to hyperoxia (200, 300%) increased
with leech size and decreased with increased temperature. The survivorship time for all size classes of N. obscura and E. punctata greatly exceeded the maximum
recorded duration (1 – 2 days) of hyperoxia in the field
during the spring. However, during the summer (20°C),
if hyperoxiain the 200 – 300% range persisted for more
than 20 days, medium and large N. obscura would
show higher mortality than comparable sizes of E.
punctata (Davies and Gates 1991a, b).
With very low or undetectable levels of superoxide
dismutase in N. obscura, Singhal and Davies (1987)
concluded that N. obscura had two possible defenses
against hyperoxia. These were to move away from areas of hyperoxia or, as recorded by Singhal and Davies
(1987), to secrete large amounts of mucus over the epidermis, which reduces oxygen diffusion into the body
and thereby the effects of oxygen toxicity. Davies and
Everett (1977) and Davies et al. (1977) recorded intraand interspecific differences in the timing of seasonal
movements between lentic microhabitats of N. obscura
and E. punctata. The results from these investigations
on hyperoxia show that small leeches are highly intolerant of hyperoxia, especially at higher temperatures,
473
and suggest that their seasonally earlier movement to
deeper water is related to avoidance of potentially
lethal oxygen conditions. Similarly, the presence of
only large leeches in the macrophyte zone, where hyperoxia is more common and more persistent, is compatible with the greater tolerance of the large size
classes to hyperoxic conditions.
During winter, leeches frequently have to rely on energy stored during periods favorable to energy acquisition although energetic demands can be mitigated by decreased activity. During the winter N. obscura exhibits
degrowth (Dratnal and Davies, 1990) and posthatchlingsized individuals that have over wintered are different,
from similarly sized individuals hatched in the summer
with respect to ecophysiological process, life-history and
fitness characteristics (Dratnal and Davies, 1990). The
cryoprotective glycerol concentration in winter leeches
are significantly higher (Reddy and Davies, 1993b), but
they have had only half the total lipid concentration energy reserves of similarly sized individuals hatched in the
summer. Under optimal feeding conditions, lipid depletion at maturity is fully compensated for in over wintered
leeches through increased food ingestion, higher absorption efficiency, and the allocation of more energy to lipid
accumulation than other growth functions (Reddy et al.,
1992). At, or just prior to sexual maturity, both winter
and summer leeches show decreases in total protein,
glycogen, and total lipid as a result of the increased energetic demand of gametogenesis and maturation.
Ultrastructural investigations of the neurons of N.
obscura exposed to anoxia and hyperoxia (Singhal et
al., 1988) showed a marked reduction in numbers of
mitochondria, ribosomes, and neurotransmitter vesicles
with swollen and fragmented cristae. These changes are
more severe in anoxia than hyperoxia.
In most lakes and rivers inhabited by leeches, ionic
content and total dissolved salts (TDS) in the water
show little temporal variability. However, significant
geographic variability occurs between lakes and rivers
primarily related to the edaphic features of the underlying geological formations. It has been shown that the
distribution of leeches is, at least in part, related to
TDS of the habitat waters (Mann, 1968; Herrmann,
1970a, b; Scudder and Reynoldson and Davies, 1976).
Similarly, in Europe, Mann (1955) showed that the
composition of the leech fauna in lentic habitats was a
function of water hardness. It has been demonstrated
that the relative concentrations of ions in the medium
modify the mortality of aquatic animals (Beadle, 1939;
Croghan, 1958; Linton et al., 1983a, b).
The blood of leeches is relatively concentrated
compared to oligochaetes and is hyperosmotic to the
water in which they live. Nephelopsis obscura and
Helobdella stagnalis were able to maintain themselves
474
Ronald W. Davies and Fredric R. Govedich
hyperosmotically in a medium between 15.6 and 59.5
mOsm / L and 47 and 112.7 mOsm / L, respectively, but
were conformers at higher medium concentrations.
Theromyzon trizonare was a regulator in both hypoand hyperosmotic media. Leeches are also capable of
regulating body volume when exposed to varying salinities (Rosca, 1950; Madanmohanrao, 1960; Smiley and
Sawyer, 1976; Reynoldson and Davies, 1980; Linton et
al. 1982). Nephelopsis obscura was able to maintain its
weight in medium from 100 – 207 mosmol / L while
E. punctata could only regulate between 58 and 200
mOsm / L. Weight regulation and volume regulation are
probably not regulated per se, although these variables
change in a predictable fashion with change in the osmotic concentration of the environment. It is more
likely that leeches regulate the osmotic pressure of the
body fluids; hence, with influxes and effluxes of water
and salts, weight and volume change.
Using a three-way factorial experimental design,
the effect of water temperature, ionic content, and TDS
on the mortality and reproduction of N. obscura and
E. punctata were examined (Linton et al., 1983a, b). A
strong interaction among temperature, ionic content,
and TDS was observed on the mortality of E. punctata,
but N. obscura showed uniformly low mortality unaffected by temperature, ionic content, or TDS. For both
species, temperature showed the greatest influence on
cocoon production and ionic content showed the least.
Cocoon production by E. punctata was reduced in
waters with low TDS and the proportion of nonviable
cocoons was higher in low TDS than in high TDS.
IV. ECOLOGY
A. Diversity
Leeches are represented in North America by four
families. Glossiphoniidae are either predators of
macroinvertebrates or temporary ectoparasites of freshwater fish, turtles, amphibians, or water birds; Piscicolidae are parasites of fishes as well as some Crustacea;
Erpobdellidae are primarily predators of macroinvertebrates and zooplankton; and, Hirudinidae are either
predators of macroinvertebrates (e.g., oligochaetes,
snails) or bloodsucking ectoparasites of freshwater and
terrestrial Amphibia and mammals. Sanguivorous
(bloodsucking) leeches spend a relatively short period
of time taking a blood meal on the host and are more
frequently found free living in the benthos. For this reason, host specificity is not often a very useful feature
for identification of leeches.
While the majority of leeches parasitic on fish
belong to the family Piscicolidae, two species of Glossi-
phoniidae (Actinobdella inequiannulata and Placobdella pediculata) show a strong preference for fish
hosts. Placobdella pediculata recorded mainly from the
midwestern United States has a high degree of host
specificity for the freshwater drum Apolodinotus grunniens (Bur, 1994). However, other species of glossiphoniids (e.g., P. montifera and Desserobdella phalera)
are also occasionally found on fishes. Members of the
Erpobdellidae and Hirudinidae have been recorded on
fishes, but it is doubtful that any species in these families regularly feeds on these hosts. All leeches require a
firm substrate for attachment and are capable of feeding on body fluids. If a healthy or injured fish can provide either of these requirements, freshwater leeches
will attach themselves. Parasitic leeches generally attach periodically to the fishes, take a blood meal, and
then move back into the benthos. Acanthobdellids feed
similarly on coldwater fishes (Bur, 1994).
B. Foraging Relationships
Leeches are either predators, consuming macroinvertebrates in the benthos and plankton or ectoparasitic sanguivores, feeding on vertebrates. The effects of
predatory fish on benthic communities have been
widely studied some showing weak or no effects
whereas other studies have shown strong effects. Dahl
and Greenberg (1997) compared prey consumption by
brown trout (Salmo trutta) and the leech Erpobdella
octoculata and showed that while one leech consumes
much less than one trout, the role of leeches in structuring benthic communities is very similar.
Leeches and acanthobdellids are primarily fluid
feeders; because, even though some predators ingest
whole prey, they quickly evacuate the hard skeletal
parts through the mouth or anus after extracting body
fluids. Sanguivory occurs in the Hirudinidae, which
have three-toothed jaws used to bite and penetrate the
host’s skin and in the Piscicolidae, and some Glossiphoniidae, which in contrast, lack jaws and penetrate
the host with a proboscis. The acanthobdellid similarly
has a short proboscislike specialization of the anterior
foregut.
Theromyzon are sanguivorous glossiphoniids feeding on waterfowl and require a minimum of three blood
meals to reach maturity (Davies, 1984; Wilkialis and
Davies, 1980b). In comparison, Placobdella papillifera
and Haementaria ghilianii require a minimum of four
(Davies and Wilkialis, 1982; Sawyer et al., 1981). The
glossiphoniids generally appear to require significantly
fewer blood meals than Hirudo medicinalis, which
takes 10 or more meals to reach maturity (Blair, 1927;
Pütter 1907, 1908), although in the laboratory at 26°C,
only 4 – 5 blood meals are required. For T. trizonare,
13. Annelida: Euhirudinea and Acanthobdellidae
475
FIGURE 13 Changes in mean weight (0 – 0) and the percentage of survival (0 – 0)
for populations of Theromyzon rude exposed to different water temperatures and
feeding regimes: (a) fed once, maintained at 20°C; (b) fed once, maintained at
5°C; (c) fed twice, maintained at 20°C; (d) fed twice, maintained at 5°C; (e) fed
three times, maintained at 20°C; (f) immature (14.7 mg) collected from the field,
fed once, maintained at 20°C; and (g) immature (14.7 mg) collected from the field,
fed once, maintained at 5°C.
over 80% of the population takes three meals in the
first 6 months after hatching (Fig. 13). The remainder of
the population overwinters after two meals and take the
third in the spring so that all the population reproduces
approximately 12 months after hatching. Some individuals that overwinter after three meals decline in weight
before spring and require a fourth meal before reproduction commences. When ingesting blood, pathogens
such as syphilis, erypsipelas, tetanus, cholera, hepatitis
B and human immune deficiency virus (HIV) enter the
gut and can survive for long periods (Nehili et al.,
1994). However, as these pathogens do not penetrate
the unicellular leech salivary glands, direct transmission
to another host is unlikely.
Predatory leeches either suck fluids with a proboscis (Glossiphoniidae) or have a suctorial mouth
(Erpobdellidae and Hirudinidae). The range of prey
species is normally quite diverse and changes seasonally
with prey availability. In some instances, prey selection
varies intraspecifically with body size and / or age class
(Davies et al., 1978, 1981, 1988; Wrona et al., 1979,
1981). Because leeches either do not ingest identifiable
hard parts or quickly evacuate them, the only accurate
and efficient method of examining feeding is with serologic techniques. Antisera against prey are produced
(Davies, 1969) and then used for a serologic test of the
gut contents.
The feeding of Nephelopsis obscura and Erpobdella punctata was investigated using specific rabbit antisera against Cladocera / Copepoda, Chironomidae,
Oligochaeta, Amphipoda, and Gastropoda. Apart from
the absence of Gastropoda in the diet of E. punctata,
there were no significant differences at the species level
in prey utilization, niche breadth, and evenness (Fig.
14). However, temporal differences in prey utilization
were evident, and both intra- and interspecific resource
476
Ronald W. Davies and Fredric R. Govedich
FIGURE 14 Frequency of prey utilization histograms (%) for Nephelopsis obscura and
Erpobdella punctata.
partitioning occurred as a function of different weight
class utilization of prey (Davies et al., 1981; Martin
et al., 1994c). Erpobdella punctata has also been
recorded as a phoront (Khan and Frick, 1997) on salamanders, with the salamander providing the leech with
food particles from its feeding. Similarly, temporal differences in feeding and intraspecific weight class differences in prey utilization were found in sympatric Glossiphonia complanata and Helobdella stagnalis (Wrona
et al., 1981) (Fig. 15). The size of both predators significantly affected feeding success on prey (Martin et al.,
1994a, b).
In freshwater ecosystems, benthic species have a
highly heterogeneous spatial and temporal distribution
and invertebrate predators, including leeches, experience variable interfeeding intervals. Smith and Davies
(1996a,b) measured the acquisition of energy and its
allocation to components of bioenergetic balance
throughout the life cycle in three groups of N. obscura
fed at low, medium and high frequency. As feeding frequency increased, the total amounts of energy ingested,
feces plus mucus produced, somatic and reproductive
growth, energy storage (total lipid), and respiration all
FIGURE 15
Frequency of prey utilization histograms (%) for
Glossiphonia complanata and Helobdella stagnalis (C / C, Cladocera /
Copepoda; C, Chironomidae; O, Oligochaeta; A, Amphipoda; G,
Gastropoda).
13. Annelida: Euhirudinea and Acanthobdellidae
increased. In the medium and high frequency feeding
treatments, the relative proportion of growth energy allocated to storage was constant, i.e. the ratio of reserves to structural tissue remains the same.
While most predatory leech species utilize a variety
of prey, Motobdella montezuma is an exception. The
pelagic endemic amphipod Hyalella montezuma comprises nearly 90% of the diet of the endemic M. montezuma in the thermally constant environment of Montezuma Well (Arizona), even though numerous other
potential prey are abundant throughout the year (Blinn
et al., 1987; Davies et al., 1988). This restricted diet
was confirmed by both gut content and serological
analyses. The highly stable conditions in Montezuma
Well have contributed to the very close relationship between the endemic predator and prey. Using a bioenergetic model McLoughlin et al. (1999) showed that the
specialized foraging utilized by M. montezuma is more
efficient to the benthic opportunistic foraging strategy
typical of most other erpobdellids. However, rather
than being an obligate forager on H. montezuma,
which is abundant during most of the year, Motobdella
montezuma exploits other prey types during the winter,
thereby achieving an overall higher rate of energy gain.
C. Reproduction and Life History
All leeches are hermaphrodites showing protandry
or cosexuality (Davies and Singhal, 1988), with reciprocal cross-fertilization as the general rule. Fertilization, which is internal, is accomplished in the majority,
of the Glossiphoniidae and all of the Piscicolidae and
Erpobdellidae by attaching a spermatophore to the
body of the partner. The spermatozoa penetrate the
body wall and make their way to the ovisacs via the
coelomic sinuses. The clitellar region is the most frequent site for the deposition of spermatophores, but
they can be attached to any part of the body of the
partner. In some species of Piscicolidae, there is a specialized region for the reception of spermatophores;
fertilization is affected only by spermatophores deposited there. In the Hirudinidae, reciprocal internal
fertilization is brought about by the insertion of an
eversible penis into the vagina of the partner.
Once fertilization occurs, the eggs are deposited
into a cocoon secreted by the clitellum. It has been
widely assumed that the presence of a visible clitellum
closely parallels maturation of the female reproductive
system, and in nonreproductive leeches the clitellar
glands are barely distinguishable from the epithelial
cells (Fernandez et al., 1992). The clitellum is especially
prominent in Erpobdellidae and has been used by numerous authors to determine sexual maturity. Biernacka and Davies (1995) showed that sexual maturity
477
of Nephelopsis obscura cannot be judged by the presence or absence of an externally visible clitellum. Although a high proportion of the population exhibited a
visible clitellum at some time, it was not persistent
(lasting no longer than 7 days) and thus could easily be
missed in studies with weekly or longer sampling periods. Feeding regime affects the timing of the clitellum
development but, regardless of feeding regime, all the
leeches exhibited two periods with a visible clitellum.
The first appearance of a clitellum did not coincide
with either spermatogenic or oogenic maturity, but at
the second appearance of a clitellum fully mature ova
were present. The presence of a visible clitellum is thus
not a good indicator of maturity in Nephelopsis obscura and probably not a good indicator in most or all
species of Erpobdellidae. The presence of a spermatophore is also not a good indicator of maturity.
Singhal et al. (1985) found only 5% of mature N. obscura collected from the field had observable spermatophores, and Biernacka and Davies (1995) found
that only 4% of the mature size range animals had a
spermatophore attached.
There can be a considerable delay between copulation and cocoon deposition, e.g. in field populations of
Helobdella stagnalis copulation occurs in the fall and
cocoon deposition takes place in the spring (Davies and
Reynoldson, 1976). Erpobdellid cocoons are thick
walled, oval, and attached to a firm substrate (stone,
leaf, stem, wood). The cocoons of Piscicolidae and
Hirudinidae are only loosely attached to the substrate
and are usually spherical. In species of Hirudinids depositing their cocoons in moist habitats out of water,
the outer wall of the cocoon is spongy, which is
thought to reduce water loss. The cocoons of Glossiphoniidae are very thin-walled (sometimes called egg
cases) and are either deposited on the substrate and immediately covered by the body of the parent or are attached to the ventral surface of the parent. In both
cases, the hatchlings are attached to the ventral body
wall and are carried around by the parent for a long
period, which for Theromyzon tessulatum can be 5
months and for T. trizonare, 1 month (Wilkialis and
Davies, 1980a). Glossiphonia complanata adults have
been found to provide their young with nutrients that
are passed through the body wall of the parent into the
posterior sucker of the young (De Eguileor et al.,
1994). Reductions in body weight and size have been
recorded for brooding glossiphoniids; however, these
losses have been attributed to reduced feeding potentials and to the energy expended ventilating the young
(Calow and Riley, 1982; Milne and Calow, 1990). It is
possible that other species which brood attached young
may provide nutrients to the developing young; however, further studies need to be conducted on a variety
478
Ronald W. Davies and Fredric R. Govedich
of glossiphoniid leech species. A comparison on reproductive output for glossiphoniid that brood eggs and
young, and for erpobdellids, that leave their eggs in an
encapsulated cocoon, shows that brooding is not metabolically more expensive than encapsulation. There are
no extra energy costs incurred in terms of carrying or
ventilating broods, but weight loss due to reduction in
feeding does occur. These costs appear to be energetically similar to the cost of encapsulation for erpobdellids, and it is suggested that brooding and encapsulation represent alternative evolutionary paths for brood
protection.
The life cycles of all leeches and acanthobdellids
consist of egg, juvenile, and mature hermaphrodite
adult, which reproduces and produces more eggs.
Adults of predatory species usually reproduce only
once before death (i.e., semelparity), although some
sanguivorous species reproduce several times over a
few years (i.e., iteroparity). Semelparity and iteroparity
are generally considered genetically different reproductive strategies determined through evolutionary
processes. Studies on the reproductive biology of
leeches have shown considerable variation in life history with annual or biennial life cycles in predatory
species. Some sanguivorous species (e.g., T. trizonare)
live for several years with the duration of the life cycle
dependent on the interval between blood meals
(Davies, 1984). Baird et al. (1986, 1987b) showed that
changes in water temperature, size at reproduction, energy loss during reproduction, and postreproductive
feeding of the erpobdellid Nephelopsis obscura significantly affected postreproductive mortality. Although
genetically iteroparous, N. obscura is like most predatory leeches and some sanguivorous species in being almost always semelparous in the field because of high
postreproductive mortality. Thus, it is quite possible
that the range of life histories reported for various
species of leeches falls within the range of environmentally induced variability. Indeed, it is conceivable that
all leeches are genetically iteroparous but phenotypically exhibit semelparity under most conditions. This
flexibility would ensure the long-term persistence of
genotypes in a variable environment, where a more
rigid strategy might have a low or zero fitness in some
years. Freshwater ecosystems in North America are
highly variable in terms of both abiotic (experience
oxygen concentration and temperature) and biotic
(prey availability) environmental parameters, and Bulmer (1985) suggested that iteroparity is found in variable environments as a bet-hedging or risk-avoidance
strategy.
The life cycle and life history of comparatively few
freshwater leech species in North America have been
examined. In Alberta, two generations of N. obscura
FIGURE 16 Life cycles of spring and summer hatching Nephelopsis
obscura in Alberta, Canada.
are produced annually, one in the spring and one in the
late summer. The spring generation is the progeny of
the heavier individuals from the spring generation of the
previous year and from the late-summer generation produced two years previously. The late-summer generation is produced by portions of the previous year’s
spring and late summer generations. Thus, each generation produces young after either 12 and 15 months or
12 and 19 months, although each individual reproduced only once (Davies and Everett, 1977) (Fig. 16).
Reddy et al. (1992) showed that N. obscura which had
overwintered differed physiologically, throughout their
life history, from individuals which had not. The first
generation of N. obscura in early summer is the progeny of the heavier individuals from the early summer
generation of the previous year and the second late
summer generation is produced by portions of the previous year’s early and late summer generations. The late
summer generation has a faster growth rate, higher lipid
storage, higher proportion of energy available for
growth and metabolism, and earlier valuation of larger
maximum size than the early summer generation. However, there were no differences in consumption of prey
or respiration between the two generations (Qian and
Davies, 1994). As there are no significant genetic differences between the early and late summer generations,
the differences in physiological ecology and life history
traits are attributed to differences in prehistory of the
parents. In Minnesota, N. obscura delays its age of reproduction to two years and in so doing attains a larger
13. Annelida: Euhirudinea and Acanthobdellidae
body size (Peterson, 1982, 1983). Erpobdella punctata
is usually subdominant to N. obscura and reproduces
after one year. When E. punctata is dominant, it has a
more complex cycle with reproduction after one or two
years, at a larger size, over a shorter season (Davies
et al., 1977). Helobdella stagnalis can produce either
one or two generations per year depending on the water
temperature regime (Davies and Reynoldson, 1976). In
warmer conditions, the overwintering population reproduces in the spring and dies on completion of brooding.
The spring generation grows rapidly and a second generation of young is produced in the summer, which
forms the overwintering population. In colder temperature regimes, members of the overwintering population
reproduce in the late spring / early summer and die after
brooding. After summer growth, this generation forms
the next overwintering population. Davies (1978)
showed that populations of H. stagnalis from cold water regimes are able to produce two generations per year
in appropriate conditions.
Several environmental factors have been implicated
in affecting the reproduction of leeches, including food
availability and water temperature. Water temperature
regime can affect the relative reproductive success and
growth of species (Davies and Reynoldson, 1976;
Davies, 1978; Wrona et al., 1987), as well as depth distribution and seasonal movement (Gates and Davies,
1987). A bioenergetics simulation model of the growth
and life history of N. obscura (Linton and Davies,
1987) showed that its growth is more sensitive to prey
variation between years than to temperature variation.
Davies and Singhal (1988) showed that, in the field, N.
obscura exhibits sequential protandry in the first bout
of gametogenesis and simultaneous hermaphroditism in
the second. In the laboratory maintained under constant conditions, oogenesis initially follows spermatogenesis but mature ova first appeared shortly after
Stage 4 of spermatogenesis with the final stages of spermatogenesis lagging behind oogenesis (sequential
protogyny). However, for about 3 weeks mature spermatozoa and ova co-occur, i.e., simultaneous hermaphroditism occurs. This is quite different from what occurs in the field. Clearly, N. obscura shows plasticity in
the phenology of the development of hermaphroditism,
the differences between the field and laboratory resulting from the absence of an environmental cue. This
was subsequently shown to be a period of low temperature (3 – 5°C) for a minimum of 7 days between stages
3 and 4 of spermatogenesis.
Kalarani et al. (1995) demonstrated the presence of
ecdysone in N. obscura and showed correlations between ecdysone concentrations and gonad maturation
and between ecdysone concentration and utilization of
energy reserves during gametogenesis. Responses to
479
ecdysone varied with both the ecdysone concentration
and the duration of exposure, indicating the importance of the sequential changes in ecdysone to the target tissues. Hormonal control of reproduction in
leeches using brain homogenates has been shown by a
number of authors ( Kulkarni et al., 1980; Hagadorn
1969; Webb, 1980; Webb and Omar, 1981).
The life cycle of Acanthobdella peledina is basically annual with young immature individuals first occurring on their host in June. These individuals leave
the fish in October on reaching sexual maturity. Reproduction with egg and cocoon production occur off the
fish and have not been observed (Holmquist, 1974).
D. Dispersal
The dispersal of freshwater leeches and acanthobdellids is generally assumed to be the result of passive
transfer from one water body to another by other animals. This seems probable for parasitic sanguivorous
species, but less probable for the majority of freshwater
leeches which are predators of benthic invertebrates.
However, some examples have been recorded with
Marrinmeyeria lucida transported by birds (Daborn,
1976), and Khan and Frick (1997) showed that a high
proportion of salamanders (Ambystoma maculosum) in
South Carolina served as phoretic hosts to Erpobdella
punctata. Platt et al. (1993) similarly recorded Helobdella stagnalis, previously found on fish, phoretic on
Ambystoma tigrinum in Indiana.
Davies et al. (1982b) examined passive transfer by
ducks of two parasitic species (Theromyzon trizonare
and Placobdella papillifera) and two predatory species
(Helobdella stagnalis and Nephelopsis obscura).
Theromyzon trizonare adults were transferred both in
the nares, while taking a blood meal, and on the duck’s
body under the feathers. Placobdella papillifera were
also conveyed on the body of the duck, but adult H.
stagnalis and N. obscura were not transported. Adult
leeches ingested by ducks were not recovered from the
duck feces, but viable N. obscura cocoons were recovered from the feces of fed ducks.
Passive dispersal by wind has never been recorded,
but the transport of cocoons attached to macrophytes
is possible. Davies (1979) showed that Helobdella stagnalis, H. triserialis, Glossiphonia complanata, and
Mooreobdella fervida were dispersed to Anticosti
Island by sea currents from the Quebec north shore of
the Gulf of St. Lawrence. Dispersal by currents in large
lakes or through rivers is also highly probable, and
leeches have been recorded in the drift (Elliott, 1973).
Accidental transport of leeches by humans has not
yet been shown in North America, but has been
demonstrated in other areas of the world (Sawyer,
480
Ronald W. Davies and Fredric R. Govedich
1986). Active directional dispersion up rivers was
recorded for Percymoorensis marmorata by Richardson (1942) and for E. punctata by Sawyer (1970).
Erpobdella punctata has been found to be a phoront
on the salamander Ambystoma maculatum, and it is
likely that as the salamanders move from pond to pond
the leeches are transported (Khan and Frick, 1997).
Passive dispersal, while unlikely, does appear to be
the mechanism by which leeches colonize new ecosystems. The passive transport of individuals between systems can be estimated using genetic markers and the
level of genetic similarity between populations can be
used to estimate the amount of migration between populations. In a recent study of Erpobdella punctata, the
estimated migration rates between populations were
found to be extremely low, regardless of the distance
between neighboring populations. However, some populations are more closely related to each other than to
others indicating the potential for either migrants or
founding individuals to be transported between ecosystems (Govedich et al., 1999).
Differences in intra- and inter-population genetic
structure is primarily determined by gene flow with
poorly dispersing species exhibiting high inter- and low
intra-population variation. In general, there is greater
genetic divergence with increased physical isolation restricting gene flow. Qian and Davies (1996) determined
whether genetic differences among three populations of
Nephelopsis obscura were the result of differential dispersal or environmental heterogeneity. Two of the populations examined were from sites in Alberta, Canada,
geographically close (80 km) to each other while the
third population in Utah, was geographically distant
(1200 km) from the other sites. Despite the high probability of dispersal between the close sites and the lower
probability of dispersal between these two sites and the
distant site, one of the sites in Alberta differed at only
one locus from the Utah population and both these populations differed from the other Alberta population at
five loci. These genetic differences were correlated with
the dissolved oxygen regimes acting as a selective force
rather than geographic distance between populations.
E. Predation
In North America, predators of leeches include
fish, birds, garter snakes, newts, and salamanders
(Pearse, 1932; Bartonek and Hickey, 1969a; Bartonek
and Murthy, 1970; Bartonek and Trauger, 1975; Bartonek, 1972; Able, 1976; Arnold, 1981; Davies et al.,
1982b; Kephart, 1982; Kephart and Arnold, 1982), as
well as insects, gastropods, and amphipods (Hobbs and
Figueroa, 1958; Pritchard, 1964; Sawyer, 1970; Anderson and Raasveldt, 1974; Blinn et al., 1988). Young
and Spelling (1986), Spelling and Young (1987) and
Young (1987), reviewing predation on three lentic
leech species in Great Britain, recorded a similar range
of predators as well as interspecific leech predation and
cannibalism.
Leeches lack hard body parts and are thus difficult,
if not impossible, to identify from the gut contents of
predators. Serology is the most appropriate method for
analyzing predation on soft-bodied animals (Davies,
1969), but this approach requires prior knowledge of
the range and size of potential predators to be collected
from the field and tested. In the laboratory, Cywinska
and Davies (1989) found that four species of dytiscids,
one species of Odonata, two species of hemipterans,
and two species of amphipods fed on one or more size
classes of N. obscura. In Great Britain, Young and
Spelling (1986) also noted that species within these
taxa fed on E. octoculata, Glossiphonia complanata,
and Helobdella stagnalis in addition to triclads, trichopterans, and megalopterans. It would thus appear
that larval and adult coleopterans and nymphal
odonates and hemipterans are potentially voracious
predators of leeches throughout the Holarctic.
In experiments with predation on cocoons, Cywinska and Davies (1989) recorded two methods of cocoon consumption; either the embryos and eggs within
the cocoons were fed upon, by piercing holes through
the cocoon wall (e.g., Coleoptera larvae and adult
Hemiptera), or else the whole cocoon (sometimes, plus
attached plant leaf) was consumed (e.g., amphipods
and adult Coleoptera). Predation has also been
recorded by gastropods on E. punctata cocoons
(Sawyer, 1970) and by a mite on Haemopsis sanguisuga cocoons (Bennike, 1943). The only known vertebrate predators of leech cocoons are salamanders
(Pearse, 1932) and waterfowl (Davies et al., 1982b).
Predation rates on N. obscura in the laboratory
were inversely related to size, with individuals 30 mg
being consumed only at very low rates by certain
coleopterans. Young and Spelling (1986) found a similar trend of increased consumption of leeches by most
predators with declining leech weight. This can be explained by the inability of some potential predators to
capture and overcome the more powerful larger
leeches, rapid satiation of predators with larger prey, or
the ability of larger leeches to produce more mucus and
so impede the movements of the potential predators.
A caveat of these feeding experiments was that
predators were starved, had no alternative food available, and had no choice in size of N. obscura prey. In
such conditions, the results of the experiments probably showed the maximum potential predation rates and
suggested that predation on cocoons and size classes
10 mg might be substantial. Considering the very
13. Annelida: Euhirudinea and Acanthobdellidae
high densities of potential predators in many lakes and
ponds and the ability of some predators to consume
large numbers of leeches and / or their cocoons, the effects of predation in the field are potentially significant
in macrophyte zones.
The rapid decline in density of hatchling and small
( 10 mg) N. obscura, observed in the field by Davies
and Everett (1977), could be explained by predation in
the shallow macrophyte zones where cocoons are deposited and hatch. Placement of cocoons by M. montezuma on deeply submerged Potamogeton stems at
least 3.5 m below the water surface at the interface of
the littoral-pelagic zone in Montezuma Well may be a
strategy to reduce predation (Davies et al., 1988).
Predation in the water column is almost entirely restricted to fish, amphibia, and birds. The presence of
large numbers of leeches in the gut contents of fish
(particularly from rivers) and the commercial development of leeches for bait in some areas indicate predation of fish can be at least occasionally heavy.
F. Behavior
Sawyer (1981) suggested that the behavior of
leeches and acanthobdellids could be divided into a
number of elementary responses: crawling (vermiform,
inchworm); swimming; shortening (fast, slow); searching (head movement, body waving); and an alert posture when the body is extended and held motionless for
some time. However, species differ markedly in the frequency of expression of these different responses. Behavior of leeches can be modified by size (age) and
physiological state (starved or fed, and mature or immature) as well as by extrinsic environmental parameters monitored by eyes, sensillae, chemoreceptors, and
mechanoreceptors (T-cells).
The foraging behaviors of sanguivores and predators are usually different. Sanguivorous leeches in the
proximity of a potential host respond to changes in illumination (shadows), vibrations in the water, chemical
stimuli, and possibly differences in temperature between the host and the surrounding water. Compared
to predatory leeches, the sanguivores are less negatively
phototactic and nocturnal, especially when hungry.
Theromyzon tessulatum, T. trizonare, Piscicola geometra, and Calliobdella vivida, when hungry, have all
been found in the open water ready to attach to a host
(Herter, 1929a, b; Meyer and Moore, 1954; Sawyer
and Hammond, 1973). Hirudo medicinalis and
Theromyzon tessulatum are stimulated to attach to and
bite hosts with temperatures of 37 – 40°C (Dickinson
and Lent, 1984), and Placobdella costata is attracted
from as far as 15 cm to hosts with temperatures of
33 – 35°C (Mannsfeld, 1934). Haementaria ghilianii
481
and Hirudo medicinalis are both sensitive to water disturbances (traveling surface waves) (Sawyer, 1981;
Young et al., 1981), and it is likely that most sanguivores respond in a similar manner.
Water disturbance is frequently the first indication
of the presence of a potential host; the leech moves toward the disturbance using cues detected by its sensilla
(Friesen and Dedwylder, 1978; Friesen, 1981; Derosa
and Friesen 1981). Theromyzon tessulatum and T. trizonare respond to water disturbance in a similar manner, but adult brooding T. tessulatum are unique in that
they move toward a potential host even though they
never take a blood meal. If the parent T. tessulatum locates a host, it attaches to it and the brooding young
leave to seek their first blood meal (Wilkialis and
Davies, 1980a).
Although movements toward a potential host by
sanguivorous leeches are not specific, the next step of
determining whether or not the potential host is an appropriate source of blood involves some selectivity. After attachment to the potential host with the posterior
sucker, the leech makes a number of searching movements with the anterior portion of the body. If the host
is not appropriate, the leech detaches. Attachment to a
host does not always occur at an appropriate location.
For example, T. trizonare primarily feed in the nares;
but if one attaches to the feathers rather than the beak
it will remain on the bird for up to 30 min, moving
about in search of a suitable location to feed. Host
recognition by sanguivores presumably results from
chemoreception and / or mechanoreception.
Predatory leeches are generally more nocturnal and
negatively phototactic than sanguivores (Elliott, 1973;
Anholt and Davies, 1987; Davies and Kassera, 1989;
Davies et al., 1996). Angstadt and Moore (1997)
demonstrated that Nephelopsis obscura swimming is
entrained to a 12-h light / 12-h dark cycle and that a
circadian rhythm of swimming persists in constant
darkness. Davies et al. (1996) tested the hypotheses
that foraging behavior responses such as numbers of
meals consumed, the number of encounters with prey
and feeding time as well as activity (swimming) were
higher in the dark, and increased with the size of N.
obscura and the length of the starvation period. While
swimming activity was longer in the dark, not all the
foraging behavior variables followed this pattern.
There was an inverse relationship between swimming
time and the number of meals consumed and the poor
correspondence between swimming time and the number of prey encounters indicates that swimming is not
always directly associated with increased foraging and
is sometimes related to movements between microhabitats. Thus, increased nocturnal activity is not always
indicative of increased foraging activity.
482
Ronald W. Davies and Fredric R. Govedich
Anholt (1986) showed that N. obscura reduced its
foraging activity when satiated and this was confirmed
through direct observations of encounters and feeding
rates as a function of satiation. Increasing satiation in
N. obscura produces a threshold level above which
prey with natural escape responses cannot be successfully captured. Only a small fraction of encountered
tubificids were ingested by starved N. obscura when
the prey had a substrate refuge. The loss of prey capture ability with increasing satiation is a behavioral foraging constraint. When its threshold level is reached,
further successful foraging on prey with a substrate
refuge is impossible. This threshold level varies for different prey types, e.g., Anholt (1986) showed that the
tube-dwelling chironomids Chironomus riparius and
Glypotendipes paripes were superior prey types compared to tubificids in promoting leech growth, and concluded that T. tubifex could evade capture by N. obscura more efficiently than chironomids. This was
confirmed by direct observations which showed escape
times by chironomids longer than the escape times of
tubificids.
Groups of N. obscura were provided with sufficient prey with either a substrate (MS) or without a
substrate (NS) to allow asymptotic food ingestion without prey patch depletion. In both groups of leeches,
feeding was not reduced by low prey encounters, but
by the predator’s ability to capture more prey. Thus,
the significantly different feeding rates exhibited by MS
and NS leeches resulted from different factors determining the asymptote of food ingestion. In MS leeches
this was primarily the foraging constraint threshold,
but in NS group it was primarily the gut capacity determining maximal feeding potential (Dratnal et al.,
1992).
As MS leeches showed a higher growth efficiency,
the lower food acquisition was compensated for by allocating a greater fraction of energy to growth and / or
by higher energy absorption efficiency, a result of
longer food retention time in a less packed alimentary
canal. Thus, the foraging constraint significantly affected the internal status of N. obscura with a feedback
system between the two. Foraging results in increasing
satiation, which reduces and eventually stops foraging,
until some level of hunger (decreased satiation) is regained. Oxygen uptake (respiration) increased in fed N.
obscura suggesting an increased energetic requirement
for the processes directly or indirectly associated with
digestion with the resulting energetic trade-off producing a lower overall activity in fed leeches.
Although several modifying, behavioral factors apply to N. obscura in the field they do not preclude the
constraint on foraging by N. obscura affecting its sizespecific rate of feeding and life-history traits, especially
as chironomids and tubificids form major components
of the diet in the field (Davies et al., 1978).
The existence of mechanisms based on individual
size-dependent limitation in food ingestion are of great
importance not only for the indeterminate growth of
N. obscura but must also strongly modify size-dependent food partitioning within the predator population — an important factor in population regulation.
To behave adaptively, animals often give higher
priority to some behaviors over others. Misell et al.
(1998) looked at the effect of feeding behavior on other
behaviors, such as swimming, crawling, and shortening, in Hirudo medicinalis. The priority feeding had,
relative to other behaviors, showed that as expected
feeding occupies a high position in the hierarchy of
behaviors and dominated all the other behaviors tested.
A feeding H. medicinalis will ignore strong mechanical stimuli which normally cause vigorous escape responses to continue feeding. Although little is know
about the neural basis of feeding in H. medicinalis, the
role of serotonin has been studied. Increases in serotonin result in an increase in feeding and vice versa,
leading to the hypothesis that serotonin is effectively a
feeding hormone.
While some predators also show increased activity
in response to disturbances in the water, they are generally opportunistic, feeding on prey encountered through
either their own locomotion or the movements of the
prey. Prey detection by predator leeches has received little attention since Gee (1913) noted that Mooreobdella
microstoma responded to vibrations in the water and
chemical extracts of its prey by unidirectional swimming
and random movements of the anterior part of its body.
Davies et al. (1982a) examined the chemosensory detection of prey by N. obscura and showed that without direct tactile contact, N. obscura were unable to detect
and react to any of the common prey types tested (i.e.,
they did not show clinotaxis). Whether or not a particular prey appears in the diet of a predatory leech depends
on the probability that it will be encountered, the probability that it will be attacked once encountered, and the
probability that an attack will result in capture. Simon
and Barnes (1996) concluded that Percymoorensis
marmorata use olfactory rather than mechano sensory
cues to detect their prey (oligochaetes). Olfactory responses of the medicinal leech (Hirudo medicinalis) are
very specific although arousal and searching for prey
are primarily initiated by water movement. The response of Bdellerogatus plumbeus to its main prey Erpobdella punctata are also chemosensory (Riggs, 1980).
However, Young et al. (1995) concluded that Helobdella stagnalis, Glossiphonia complanata and Erpobdella octoculata do not depend on chemosensory cues
to detect prey and that chemosensory cues are only
13. Annelida: Euhirudinea and Acanthobdellidae
important when the leech is in contact with the prey as
shown by Davies et al. (1982a). They also concluded
that the inability of leeches to detect damaged prey in
heterogenous environments was the reason for very high
juvenile mortality.
An exception to the general rule that the diet of
predatory leeches is usually nonspecific and contains a
wide variety of prey is demonstrated by Motobdella
montezuma. Blinn et al. (1988) showed that M. montezuma forages selectively on the pelagic amphipod
Hyalella montezuma despite the presence of a wide variety of alternate prey. Motobdella montezuma discriminates between two congeneric amphipod prey by
mechanoreception, exhibiting a high response to the
acoustic vibration signals of the endemic H. montezuma but not to H. azteca. Not only does M. montezuma discriminate between species by mechanoreception, but it also selectively discriminates between size
classes, feeding principally on juveniles. Although amphipods form part of the diet of both N. obscura and
E. punctata, neither species shows a significant response to their acoustic vibration signals (Blinn and
Davies, 1989). The unusual feeding behavior exhibited
by Motobdella can be linked to the highly specialized
sensillae that ring the mouth (Fig. 17). These sensillae
are composed of cilia that are in contact with the water
where vibrations from potential prey can be detected.
The number of cilia and the size of the sensillae have
been found to be significantly greater in Motobdella
than in the related erpobdellid genera Erpobdella and
Nephelopsis (Govedich et al., 1998).
Substrate type affects leech foraging with Erpobdella octoculata foraging more effectively over cobble
substrates than over fine gravel with differences in prey
type consumed (Dahl and Greenberg, 1997). When
leeches and trout were together, trout foraging was unaffected by leeches but leech foraging was affected by
trout. Although one leech may consume much less than
one trout at field densities, leeches have similar effects
as fish in structuring lotic benthic communities (Dahl,
1998).
Some animals form temporary or permanent aggregations or groups which may reduce an individual’s
risk of predation, enhance mating or food intake, ameliorate physiological stress, coordinate and enhance dispersal or synchronize development. Leeches commonly
form aggregations in the field, especially the benthic
predatory species. Smith and Davies (1996a, b) examined the effects of different group sizes on acquisition
and allocation of energy in Nephelopsis obscura and
evaluated the physiological cost and benefits of group
living. In terms of growth, asymptotic biomass, ingestion and respiration group sizes larger than one and
less than 10 were found to be optimal — a group size
483
within the typical range of groups sizes found on natural stony shores.
Oxygen concentrations affect both the growth of
N. obscura and its feeding behavior (Davies et al.,
1992; Davies and Gates, 1991a, b). Higher rates of
feeding occurs in 100 and 300% oxygen saturation regions compared to hypoxic regimes. This is related to
the greater proportion of time spent on or in the mud
substrate and thus increased prey encounter rate. Since
hypoxia and anoxia cause high mortality at summer
temperatures, differences in vertical distribution of N.
obscura are behavioral adaptations to minimize stress.
Seasonal migrations of N. obscura from deep water to
shallow water in the spring and to deep water from
shallow water in the fall cannot be solely attributed to
temperature preferences and it is likely that avoidance
of oxygen stress also plays a role.
G. Population Regulation
On a geological time scale, only a very few freshwater ecosystems are long-lived and even on a shorter
time scale of 1 – 20 years, many lentic ecosystems are
ephemeral. In ephemeral ecosystems, leeches may not
have sufficient time to reach carrying capacity. Evidence for competition between leech species has been
presented by Davies et al. (1982c) for Nephelopsis obscura and Erpobdella punctata and by Wrona et al.
(1981) for Glossiphonia complanata and Helobdella
stagnalis. However, only the studies on competition between N. obscura and E. punctata are comprehensive
in relation to the established criteria (Reynoldson and
Bellamy, 1970; Williamson, 1972).
Reynoldson and Davies (1976, 1980) showed a
considerable overlap in the range of water conductances in which N. obscura and E. punctata can regulate their weights. The sympatric distributions of the
species, therefore, appear to be related to common
salinity tolerances. However, among sympatric populations, there is a strong inverse relationship between the
numerical abundances of N. obscura and E. punctata
in lentic ecosystems (Davies et al., 1977). Dominance
of E. punctata over N. obscura is a rare temporary
event and the switch to dominance by N. obscura is
rapid. The reproductive strategies of E. punctata vary
depending on whether it is dominant or subordinate
(Davies et al., 1977). Thus, the biological interactions
between N. obscura and E. punctata are amenable to
explanation based on competition.
The utilization of common food resources by N.
obscura and E. punctata has been demonstrated by numerous laboratory and field studies (Davies and
Everett, 1977; Davies et al., 1978, 1981, 1982c). At the
species level, there were no significant differences in the
FIGURE 17
Photomicrographs (5000 ) showing differences in the size of sensilla and the number of cilia
within sensilla of: (a) Motobdella sedonensis; (b) M. montezuma; (c) Erpobdella punctata; (d) Nephelopsis
obscura (Govedich et al., 1998).
484
13. Annelida: Euhirudinea and Acanthobdellidae
diets of either species between years and the trends in
prey utilization were similar each year (Fig. 14). Intraspecific food resource partitioning, as a function of
differential weight class utilization of prey was shown
by both species. Such resource partitioning minimizing
intraspecific competition, increases population fitness
(Giesel, 1974). The change in feeding ecology of E.
punctata with changes in numerical dominance provided further evidence for interspecific competition.
Furthermore, a significant decrease in mean niche overlap and niche breadth was found for both species when
N. obscura became dominant. The majority of criteria
established to determine interspecific competition have
been supported by the data collected on N. obscura
and E. punctata; however, the manipulation of food resources has not yet been completed. In British lakes,
semelparous leech species show very high (95 – 98%)
mortality of juveniles (Young et al., 1995). Many
young leeches are inept at capturing prey, and feeding
success increases when adult leeches are also present
suggesting that mortality of young leeches is high because of their inability to locate damaged prey in an
heterogenous environment (Dahl, 1998).
H. Ecotoxicology
Aquatic ecosystems seem to be sinks for many environmental contaminants such as heavy metals, pesticides and other synthetic or natural pollutants. The impacts of these pollutants are generally only seen when
the problems have reached critical end points such as
fish kills or algal blooms. However, more subtle effects,
such as changes in growth, reproduction, behavior,
physiology or survival of freshwater species, may act as
indicators for potential larger scale problems. In the
longer term, these subtle chronic effects play a role on
health and efficiency of freshwater ecosystems. Biomonitoring and ecotoxicological studies utilizing freshwater leeches, especially benthic predatory species,
have recently become increasingly more common.
Changes in the community are frequently assessed
by using bioindicators (species or groups of species for
which changes in abundance reflects the quality of the
environment. Leeches were used by Bendell and McNichol (1991) as indicators of acidification on 40 small
lakes, many of which had been acidified by nickel smelting works at Sudbury, Ontario, Canada. Nine species of
leeches were recorded from the 20 lakes with pH 5.5.
Several species of leech disappeared over a narrow range
of pH which suggests that acidity or a single chemical or
biological factor correlated with pH is responsible.
However, Grantham and Hann (1994) showed that lake
pH in the Experimental Lakes Area of Ontario was not
a dominant variable, describing only a small amount of
the variance in the species – environment relationship.
485
Leeches have shown superior chlorophenol bioconcentrating ability over many other aquatic organisms.
Prahacs and Hall (1996) evaluated N. obscura as a potential biomonitor of chlorinated phenolic compounds
discharged from bleach pulp mills on the Lower Fraser
River, British Columbia, Canada. They found a strong
linear correlation between water contaminant concentration (0.1 – 10.0 g / L) and bioconcentration of
chloroguaiacols. Bioconcentration was inversely related
to pH. As in situ biomonitors, Prahacs et al. (1996)
showed that and increase in chlorine dioxide substitution in leeches reduced amounts of chlorinated phenolics accumulated, with a sharp decrease observed at
chlorine dioxide levels 90%.
An alternative approach to bioindicators is the use
of biomarkers, defined as measurable variations in cellular or biochemical components or processes, structures or functions, that provides information on a stress
and its effects at the population level. The effects of exposure of adult sexually mature Nephelopsis obscura
for 4 – 24 days to 0.0 – 580 g / L cadmium showed that
the numbers of ova and spermatozoa per unit biomass
were significantly reduced with increasing cadmium
concentration and exposure time, as were the masses of
the testisacs and ovisacs (Davies et al., 1995). Exposure
of cocoons to cadmium (0.0 – 4000 g / L) had a significant effect on post hatchling survivorship (Wicklum et
al., 1997). The effects of chronic exposure to cadmium
(0.0 – 50 g / L) were examined in terms of changes in
energy acquisition and allocation (somatic growth, reproductive growth, feces plus mucus, active respiration,
resting respiration and activity time) (Wicklum and
Davies, 1996). Leeches in the high cadmium concentration group had significantly lower biomass production,
survivorship, and ingestion rate. Assimilation efficiency,
reproductive investment, resting respiration and active
respiration were unaffected by cadmium, but leeches in
the high concentration group were significantly less active and produced more mucus. Decreased ingestion
and increased mucus production in the high concentration of cadmium contributed to a lower growth rate.
Although cadmium had no effect on resting or active
respiration, there were significant effects on total respiration as leeches exposed to high cadmium doses spent
less time in active and, thus, in total respiration.
Macroinvertebrates are generally good indicators of
toxic stress in wetlands; but while phorate (a commonly
used pesticide) killed all amphipods and chironomids,
snails and leeches were tolerant and could survive heavy
doses by behavioral adaptations and / or physiological
tolerances (Dieter et al., 1996). Leeches are also not
good monitors of mercury (usually as methyl mercury)
biomagnification and are not a dietary risk to waterfowl or fish (McNicol et al., 1997). Studying polychlorinated biphenyls discharged to a lotic system in Ontario,
486
Ronald W. Davies and Fredric R. Govedich
fish and leeches occupying the top of the food web accumulated more PCBs than organisms lower in the food
web, thus indicating that biomagnification (uptake
through ingestion) through trophic transfer and not
bioconcentration is the primary mechanism governing
contamination (i.e., uptake as a chemical from the
water) (Zaranko et al., 1997).
V. COLLECTION AND REARING
A. Collection
Leech and acanthobdellid ectoparasites are sometimes found attached to hosts and are usually easily
removed. Most leech species (but not Acanthobdella
peledina) are free-living for the majority of their life
cycle and are normally attached to a firm substrate or
free swimming, as are the predatory leech species. They
can be collected with bottom samplers and dip or
plankton nets.
Free-living leeches can be sampled quantitatively
using colonization chambers, grab samplers, or timed
collections from rocks and stones. As a general rule, the
plan area of the largest leech collected, or its area of
escape response, should not exceed 5% of the sampler
area (Green, 1979). The only benthic samplers that even
approximately meet these criteria are grab samplers such
as the Ekman grab (Ekman, 1911). Another problem
faced when quantitatively sampling free-living leeches is
that they occur in both the sediment and the vegetation.
The Gerking box sampler (Gerking, 1957) is one of the
few methods of quantitatively estimating population
density, separable into these two components. This technique has, however, a number of technical problems including the difficulty of clipping macrophytes close to
the substrate, the entrainment of sediments and associated fauna in the macrophyte sample, the invasive influence of the operator, and the difficulty of integrating the
macrophyte and sediment samples because the benthic
samples are taken from inside the area from which the
macrophyte samples are collected. These problems are
overcome by using a sampler consisting of a bottom
mounted detachable grab connected to a box sampler
with levered, spring-loaded jaws for cutting macrophytes (Gates et al. 1987). The sampler is triggered from
above the water and simultaneously compartmentalizes
the soft sediments in the grab and the phytomacrofauna
in the water column above.
B. Rearing
Leeches are generally easy to maintain in the laboratory provided they are maintained under approximately
similar conditions encountered in the field. Appropriate
dissolved oxygen and temperature regimes are particularly important. Light regime is less critical, but if an
ambient light regime cannot be provided, darkened conditions are preferable. Leeches are very susceptible to
rapid changes in ionic content, temperature, and dissolved oxygen concentrations. If field conditions cannot
be provided, they should be slowly acclimated to the
new conditions (e.g., temperature changes exceeding
5°C per day will usually cause very high mortality, but a
temperature change of 1°C per day within the normal
field range will usually result in little mortality).
Predatory leech species are easily fed in the laboratory. If suitable prey at high densities are provided,
most leeches will feed avidly when hungry. For maintenance, ad libitum feeding 1 day per week is sufficient.
Most predatory species brought into the laboratory
and provided with ambient field conditions will feed,
grow, and reproduce. The viability of the cocoons produced is sometimes decreased if the parent has been
maintained in the laboratory for a long period. It is
much more difficult to raise hatchlings produced in the
laboratory through to mature size, capable of producing viable cocoons. Each species requires special conditions and there is much research to be done in this area.
Conversely, while some sanguivorous leeches are
more difficult to feed in the laboratory, reproduction,
cocoon production, and hatching are generally easier.
Some sanguivores (e.g., Placobdella papillifera) will eat
if immersed directly into blood. Many others (e.g.,
Hirudo medicinalis) will feed through a membrane if
blood at an appropriate temperature (30 – 40°C) is on
the other side. For these species, the simplest technique
is to provide the hungry sanguivore with a sausage
made from intestinal epithelium filled with warmed
blood. Other species (e.g., Theromyzon trizonare) have
not yet been induced to feed artificially and must be
provided with a live host and given the opportunity to
reach an appropriate feeding site (the nares of waterbirds for T. trizonare). However, when suitable sites are
available, cocoon production occurs readily.
VI. IDENTIFICATION
A. Preparation of Specimens
Some species can be studied live, but permanently
stained slides and serial sections are required for identification of a few species. Clearing, without permanent
staining, is often required for determining the ocular
number and arrangement.
Most leeches contract strongly when placed in cold
fixative and should be narcotized first. Before narcotizing
13. Annelida: Euhirudinea and Acanthobdellidae
the specimens, the colors of the dorsal and ventral
surfaces should be recorded, as chromatophores are
dissolved or altered by most narcotizing agents. Leeches
are generally difficult to relax properly, and any of the
following three methods are recommended:
1. Add drops of 95% methanol slowly to the
water containing the leech, gradually increasing
the concentration for about 30 min until movement ceases. When the leech is limp and no
longer responds to touch, pass it between the
fingers to straighten it and remove excess
mucous.
2. Add carbonated water or bubble in CO2 until
movement of the leech stops. Straighten the
leech out on a slide and slowly add warm 70%
ethanol and a few drops of glacial acetic acid
until it is covered.
3. Add a drop or two of 6% nembutal until
movements stop and then straighten the leech
on a slide.
For some specimens, slight flattening is occasionally desirable. This is best done between two glass slides with
weights added if necessary. After relaxation, the leeches
should be fixed for 24 h (depending on size) in 5 or
10% formalin. For histological preparations, Fleming’s
or Bouin’s fixatives should be used. Large leeches
should be injected with 70% ethanol to ensure preservation of the internal organs.
To preserve a leech, wash the fixed specimen with
deionized water to remove the formalin and then add
either 70% ethanol or 5% buffered formalin. Although
formalin preserves the colors longer, ethanol is recommended since it is safer to use and less destructive to soft
tissues. Leeches fixed in formalin or Bouin’s or Fleming’s
FIGURE 18
487
fixative can be stained in Mayer’s paracarmine, borax
carmine, or Hams’ hematoxylin for 12 – 94 h, destained
in a 1% HCl – 70% ethanol solution until the leech
epidermis is free of stain, and then neutralized in a 1%
NH4OH – 70% ethanol solution. Fast green or eosin are
routinely used as counterstains. Stained specimens
should be dehydrated in progressively higher concentrations of ethanol, cleared in methyl salicylate, and
mounted in a neutral pH mounting medium.
B. Important Features for Identification
Identification of leeches can be difficult, if not impossible, with poorly preserved specimens. External
features such as the following are used whenever possible: annulation; number of eyes and their arrangement;
presence or absence of papillae, tubercles, and pulsatile
vesicles; relative size of anterior (oral) and posterior
(anal) suckers; size and position of the mouth; and, the
relative positions of the male and female gonopores.
The only notes on coloration included in the following
keys are those known to persist after preservation. In a
few instances, reference is made to internal anatomical
features. These have been kept to a minimum and are
used only when positive identification is impossible
without them. To distinguish between Nephelopsis obscura and the Dina-Mooreobdella complex, the atrial
cornua must be examined (Fig. 18). The preserved
specimen should be pinned out with the ventral surface
up and a transverse incision made across the body
three or four annuli) posterior to the male gonopore.
Cuts should be made anteriorly up the lateral margins
of the body for about 20 annuli) and the posterior edge
of the flap lifted forward exposing the inner tissues,
which must be cleared to expose the atrium. To help
(a) Spirally curved atrial cornua of Nephelopsis obscura; and (b)
the simply curved atrial cornua of Dina / Mooreobdella.
488
Ronald W. Davies and Fredric R. Govedich
identify Desserobdella the salivary gland can be examined in specimens cleared with clove oil.
To examine the velum and jaws, the specimen
should be pinned out and a median ventral incision
made from the lower lip of the anterior sucker back far
enough for the margins to be pinned out to expose the
inner surface of the pharynx (Fig. 8). Details of the
teeth (Fig. 9) can only be seen by removal of a jaw and
making a temporary mount on a microscope slide.
The length of leeches and acanthobdellids is dependent on species, age, physiological state, and the degree
of relaxation. The measurements presented in the keys
are maximum total lengths of relaxed individuals.
C. Family Glossiphoniidae
Confusion exists in the systematics of these genera,
and many species have undergone considerable synonymy. Although some taxonomists consider that the
genus Haementeria (De Filippi, 1849) has priority over
Placobdella (Blanchard, 1893), the latter name is widely
recognized and accepted as a genus in North America
and is well represented with nine species. Lukin (1976)
recognized important differences between Glossiphonia
complanata and G. heteroclita, and suggested that the
latter species be placed in the subgenus Alboglossiphonia
elevated to generic level by Klemm (1982). There has
been considerable confusion among the species assigned
to the genera Batracobdella Viguier and Placobdella
Blanchard. Barta and Sawyer (1990) erected a new
genus Desserobdella, differentiated by the presence of
two pairs of coalesced eyes and one pair of diffuse salivary glands compared to the two pairs of pairs of compact salivary glands in Placobdella and a single pair of
eyes in Batracobdella. Placobdella species primarily specialize on reptilian and crocodilian hosts whereas species
of Desserobdella utilize amphibian or piscine hosts. Reexamination of type specimens by Barta and Sawyer
(1990) and Jones and Woo (1990) assigned Placobdella
(Batracobdella) picta, Placobdella (Batracobdella)
phalera, Batracobdella cryptobranchii and Batracobdella (Placobdella) michiganensis to the genus Desserobdella. Placobdella parasitica and P. ornata have been
confirmed as members of the genus Placobdella (Moser
and Desser, 1996), but similar re-examination of the
number and structure of the salivary glands is still
needed for P. hollensis, P. pediculata, P. montifera, P.
nuchalis, P. translucens, P. mulilineata, and P. papillifera.
The single and questionable identification of the
European species Batracobdella paludosa by
Pawlowski (1948) from Nova Scotia, Canada is the
only record of this genus from North America.
Theromyzon tessulatum is characteristically a
European species, but its presence in North America
has been well documented (Davies, 1971, 1973, 1991;
Klemm, 1982; Sawyer, 1986; Oosthuizen and Davies,
1993). The first species of Theromyzon reported from
North America, described by Baird (1869) as Glossiphonia rudis, was based on a brief nondiagnostic description. Because Moore and Meyer (1951) and Meyer
and Moore (1954) collected specimens from the same
locality (Great Bear Lake) they assumed their specimens with the male and female gonopores separated by
three annuli were Theromyzon rude. Thereafter, all
specimens of Theromyzon collected in North America
with three annuli between the gonopores have been
identified as T. rude. However, examination and dissection of the syntypes (Oosthuizen and Davies, 1992)
showed that one specimen was Placobdella ornata and
the second, which became the holotype of Glossiphonia rudis, had only two annuli separating the gonopores. Specimens with three annuli between the gonopores were subsequently shown to belong to a new
species T. trizonare and all specimens previously identified as T. rude are, in fact, T. trizonare. In North
America, there are three species with two annuli between the gonopores (Oosthuizen and Davies, 1993):
T. maculosum, T. bifarium, and T. rude. Theromyzon
maculosum, characteristically a European species, is
easily distinguished from T. bifarium and T. rude by
the presence of two independent oviducts with two female pores compared to one female pore in the latter
two species. Theromyzon bifarium is distinguished
from T. rude by the presence of a cylindrical male
atrium, paired dorso-lateral male ducts which enter the
atrium separately. Although T. sexoculatum (Moore,
1898) and T. biannulatum (Klemm, 1977) have been
described from North America with two annuli between the gonopores they are not valid species.
The genus Actinobdella was at first erroneously
placed in the family Piscicolidae by Moore (1901), who
was misled by the unusual annulation of Actinobdella
inequiannulata. Complete segments of Actinobdella are
3-annulate (a1, a2, a3), which are further divided into
six unequal annuli (b1, b2, b3, b4, b5, b6).
D. Family Piscicolidae
Although the Family Piscicolidae has been shown
to be paraphyletic (Siddall and Burreson, 1995) it has
been retained for convenience and is based on plesiomorphic characteristics and parasitism of fish.
Some workers have suggested that Piscicola
geometra and P. milneri are synonymous, although
P. milneri has 10 – 12 punctiform eyespots, whereas
P. geometra has 12 – 14 punctiform eyespots and dark
pigmented rays on the posterior sucker (Fig. 19c, d).
Klemm (1977) indicated that the number of punctiform
13. Annelida: Euhirudinea and Acanthobdellidae
489
relatively simple to combine the records for Illinobdella
with those for Myzobdella. However, if supporting
data are not forthcoming, subsuming Illinobdella with
Myzobdella will result in an irretrievable loss of information.
E. Family Hirudinidae
Richardson (1969, 1971) divided the North American Haemopisoid leeches into the genera Mollibdella,
Bdellarogatis, and Percymoorensis, rather than placing
them all in the genus Haemopis. This was accepted by
Soös (1969b), Davies (1971 – 1973), and Klemm
(1972a), and is accepted here. Sawyer and Shelley
(1976) and Sawyer (1986), for unspecified reasons, recombined them all back into the genus Haemopis, a
view also adopted by Klemm (1977, 1982, 1985).
FIGURE 19 Posterior suckers of: (a) Piscicola punctata without
oculiform spots; (b) Piscicola salmositica with 8 – 10 crescentiform
oculiform spots; Piscicola geometra with 12 – 14 oculiform spots
separated by pigmented rays; and (c) Piscicola Milnera with 10 – 12
punctiform oculiform spots on posterior sucker.
eyespots on the posterior sucker and the presence or
absence of the pigmented rays varies with the age of
the specimens. As the gonopores of Piscicola milneri
are separated by two annuli, and by three annuli in P.
geometra, they are here considered distinct species.
The genera Myzobdella and Illinobdella are very
similar (Meyer, 1940; Daniels and Sawyer, 1973), but
their relationship needs to be thoroughly investigated
before the synonymy suggested by Daniels and Sawyer
(1973) and Sawyer (1986) can be fully accepted. In the
present chapter, a conservative approach has been used.
If later evidence supports the propositions of Daniels
and Sawyer (1973) and Sawyer (1986), then it will be
F. Family Erpobdellidae
There is some disagreement as to whether Dina
and Mooreobdella are distinct genera (Moore, 1959) or
subgenera of the genus Dina (Soös 1968). The diagnostic feature is whether the vasa deferentia form preatrial
loops extending to ganglion XI (Dina) or do not have
preatrial loops (Mooreobdella). This is difficult to determine, and thus in this key, the genera (or subgenera)
are lumped together in a Dina – Mooreobdella complex, separated on external features. Although placed
in the genus Erpobdella by Davies et al. (1985), Motobdella montezuma has several morphological traits
which distinguish it from Erpobdella including posterior and crop ceca, large sensilla and the location of the
anus next to the posterior sucker. Motobdella sedonensis can readily be distinguished from M. montezuma in
having fewer papillae (10 – 14 vs 14 – 18) and in having
the papillae on the dorsal surface not extending round
the body as M. montezuma.
VII. TAXONOMIC KEYS
A. Euhirudinea and Acanthobdella
1a.
Body divided into 29 postoral segments; segments subdivided superficially into four annuli (a1, a2, b5, b6) (Fig. 2b): no anterior
sucker, mouth on ventral surface of segment III; no jaws; posterior suckering of four segments: two pairs of chaetae on five consecutive anterior segments; distal ends of chaetae bent to form hooks (Fig. 4b); chaetae absent from remainder of the body; paired ventral male gonopores: single median ventral male gonopore; (Fig. 4a) length 22 mm; ectoparasite of salmonids....................................
Acanthobdella peledina Grube, 1851
1b.
Body divided into 32 postoral segments; segments subdivided superficially into 3 – 16 annuli (Figs. 2a and 31), anterior sucker
present consists of four segments: mouth on ventral surface of anterior sucker (Fig. 7); jaws present or absent; posterior sucker
consists of seven segments; chaetae absent from entire body; median ventral unpaired male and female gonopores (Figs. 3 and 29)
..........................................................................................................................................................................................Euhirudinea
490
Ronald W. Davies and Fredric R. Govedich
B. Species of Euhirudinea
1a.
Mouth a small pore on ventral surface of anterior sucker through which a muscular pharyngeal proboscis can be protruded (Fig.
7a, b); no jaws or teeth.........................................................................................................................................................................
Rhynchobdellida ...............................................................................................................................................................................2
1b.
Mouth large, occupying the entire cavity of anterior sucker (Fig. 7c); no protrusible proboscis; jaws with teeth either present or
absent (Fig. 8).................................................................................................................Arhynchobdellida........................................3
2a(1a)
Body flattened dorso-ventrally and much wider than head (Fig. 1a) (except Placobdella montifera and P. nuchalis); body not cylindrical (except for Helobdella elongata which is subcylindrical); not differentiated into two body regions: anterior sucker ventral,
more or less fused to body and narrower than body; body never divided into anterior trachelosome and posterior urosome; eggs in
membranous cocoons and young brooded on ventral surface of parent; one, two, three or four pairs of eyes; no oculiform eye spots
on posterior sucker; segments 3-annulate (a1, a2, a3) (Fig. 2a) (except Oligobdella biannulata which is 2-annulate)...........................
Family Glossiphoniidae ......................................................................................................................................................................4
2b.
Body cylindrical and usually long and narrow; body sometimes divided into a narrow anterior trachelosome and a wider posterior
urosome (Illinobdella and Piscicolaria) (Figs. 1b, 6); anterior sucker expanded and distinct from body: zero, one, or two pairs of
eyes; pulsatile vesicles along the lateral margins present (Piscicola and Cystobranchus) or absent; seven or more annuli per segment
(except Piscicolaria reducta which is 3-annulate); oculiform eye spots sometimes present on posterior sucker (Figs. 6b, 19b – d); no
brooding of cocoons or young ...................................................................................................................................................Family
Piscicolidae.......................................................................................................................................................................................37
3a(1b)
Five pairs of eyes arranged in an arch on segments II – VI with the third and fourth pairs of eyes separated by one annulus
(Fig. 20a); body elongate (Fig. 1c); jaws with teeth either present or absent; nine or ten pairs of testisacs arranged metamerically
(Fig. 11a); pharynx short ...........................................................................................................................................................Family
Hirudinidae......................................................................................................................................................................................49
3b.
Zero, three, or four pairs of eyes in separate labial and buccal groups (Fig. 25b); body elongate (Fig. 1d); no jaws; testisacs small
and numerous (Fig. 11b); pharynx about one-third of body length .....................................................................Family Erpobdellidae
.........................................................................................................................................................................................................62
4a(2a)
Posterior sucker conspicuous with a marginal circle of 30 – 60 glands and retractile papillae, their positions being indicated dorsally
by faint radiating ridges (Fig. 21) .........................................................................................................................................................
Actinobdella .......................................................................................................................................................................................5
4b.
Posterior sucker without a marginal circle of glands or retractile papillae ..........................................................................................6
5a(4a)
Posterior sucker on short, distinct pedicel with 29 – 31 digitate processes on rim (Fig. 21); somites 3- or 6-annulate; dorsal papillae
in 1 – 5 longitudinal rows; length 22 mm ............................................................................Actinobdella inequiannulata Moore, 1901
5b.
Posterior sucker on short distinct pedicel with about 60 digitate processes on rim; somites six-annulate with b3 and b5 the largest
and most conspicuous; length 11 mm; dorsal papillae in five longitudinal rows .........................Actinobdella annectens Moore, 1906
6a(4b)
Zero, one, or two pairs of eyes; a series of paired accessory eyes sometimes present along body (Fig. 5c) ..........................................7
6b.
Three or four pairs of eyes ...............................................................................................................................................................30
7a(6a)
Mouth apical or subapical on rim of anterior sucker (Fig. 7a); zero ,one or two pairs of eyes ............................................................8
FIGURE 20
Arrangement of eyes in: (a) Hirudinidae; and (b) Erpobdellidae. FIGURE 21 Lateral view
of the posterior sucker of Actinobdella inequiannulata showing the pedicel and the retractile papillae around
the margin.
13. Annelida: Euhirudinea and Acanthobdellidae
491
7b.
Mouth within anterior sucker and clearly not on rim (Fig. 7b); one or two pair of eyes; gonopores separated by one or two annuli
(Fig. 3b) ...........................................................................................................................................................................................18
8a(7a)
Male and female gonopores united in common bursal pore; one pair of eyes well separated; body smooth without papillae; six pairs
of crop ceca; length 22 m..............................................................................................................Marvinmeyeria lucida Moore, 1954
8b.
Male and female gonopores separated by two annuli (Fig. 3b); zero or one pair of eyes (Fig. 5a) [Placobdella hollensis has several
pairs of accessory eyes on the neck (Fig. 5c)]; eyes close together or confluent (Fig. 5b) (except Placobdella montifera and P.
nuchalis which have eyes well separated); body usually papillated; seven pairs of crop ceca ................................................................
Placobdella / Oligobdella ....................................................................................................................................................................9
9a(8b)
One small pair of eyes on segment II and one pair of larger sometimes coalesced eyes; no supplementary eyes ................................10
9b.
One pair of small eyes on segment II and a larger sometimes coalesced pair of eyes on segment III followed by an indefinite number
of pairs of accessory eyes (Fig. 5c); tubercles large and rough; length 30 mm...........................Placobdella hollensis (Whitman, 1892)
10a(9a)
Anus between segments XXIII and XXIV with the 16 postanal annuli forming a slender stalk (pedicel) which bears the posterior
sucker; no papillae; length 35 mm .....................................................................................Placobdella pediculata (Hemingway, 1908)
10b.
Anus close to the posterior sucker; no pedicel ..................................................................................................................................11
11a(10b)
Margins of posterior sucker denticulate (Fig. 22) head expanded and discoid and set off from body by a narrow neck (Fig. 23) .....12
11b.
Margins of posterior sucker not denticulate .....................................................................................................................................13
12a(11a)
Dorsum with three prominent tuberculate keels or ridges (Fig. 23); length 16 mm.............................................................Placobdella
montifera Moore, 1906
12b.
Dorsum smooth; no keels or ridges; length 25 mm ......................................................Placobdella nuchalis Sawyer and Shelley, 1976
13a(11b)
Medium longitudinal row of tubercles on dorsal surface are large and conspicuous; tubercles bear several papillae giving a rough,
warty appearance; ventrum unstriped; length 40 mm.......................................................................Placobdella ornata (Verrill, 1872)
13b.
Dorsal papillae, if present, simple cones with smooth domes ...........................................................................................................14
14a(13b)
Dorsum smooth; dorsal papillae inconspicuous or absent ................................................................................................................15
14b.
Dorsal papillae small and conical but distinct ..................................................................................................................................17
15a(14a)
Dorsum smooth; no papillae or tubercles; dorsum with conspicuous white genital and anal patches; one or more medial white
patches; white bar on neck ...............................................................................................................................................................16
15b.
Dorsal papillae inconspicuous or absent; when present, papillae arranged in five longitudinal rows; ventral surface with 11 – 12
blue, brown, or green stripes; annulus a3 in middle of body without distinct cross furrow; length 65 mm ...........................................
Placobdella parasitica (Say, 1824)
FIGURE 22
Posterior sucker of Placobdella montifera showing the denticulate margins. FIGURE 23
Placobdella montifera showing the expanded discoid head and dorsum with three prominent tuberculate
dorsal ridges (keels).
492
Ronald W. Davies and Fredric R. Govedich
16a(15a)
3-annulate (Fig. 2a); posterior sucker not conspicuous; length 11 mm .............................................................Placobdella translucens
Sawyer and Shelley, 1976
16b.
2-annulate; posterior sucker large; length 7 mm ...............................................................................................Oligobdella biannulata
(Moore, 1900)
17a(14b)
Dorsum with 5 – 7 longitudinal rows of small papillae; papillae on posterior sucker; annulus a3 in middle of body with distinct cross
furrow; length 45 mm ................................................................................................................Placobdella papillifera (Verrill, 1872)
17b.
Dorsum with few or numerous small papillae in five longitudinal rows; length 50 mm ..................................Placobdella multilineata
Moore, 1953
18a(7b)
One or two pairs of eyes; if only one part of eyes present, these are close together or lobed indicating coalescence (Fig. 5a, b); gonopores separated by two annuli (Fig. 3b)............................................................................................................................................19
18b.
One pair of eyes which are well separated (Fig. 24); gonopores separated by one annulus .............................................Helobdella 23
19a(18a)
One large pair of eyes on segment IV sometimes coalesced (Fig. 5a, b); seven pairs of crop ceca; papillae absent; length 20 mm
...............................................................................................................................................Batracobdella paludosa (Carena, 1824)
19b.
One pair of eyes on somite II and a larger pair of eyes on segment III eyes sometimes confluent (Fig. 5b); one pair of diffuse salivary
glands; two annuli separating gonopores (Fig. 3b) ...............................................................................................................................
Desserobdella ...................................................................................................................................................................................20
FIGURE 24 Helobdella stagnalis (or H. california) showing the chitinous scute (nuchal plate) on the dorsal
surface and the single pair of eyes. FIGURE 25 Dorsal view of: (a) Theromyzon trizonare; (b) Glossiphonia
complanata; and (c) Alboglossiphonia heteroclita showing the arrangement of the eyes. FIGURE 26 (a)
Myzobdella lugubris; and (b) Piscicolaria reducta showing the trachelosome and urosome without pulsatile
vesicles, the relative sizes of the eyes, and the weakly developed posterior sucker.
13. Annelida: Euhirudinea and Acanthobdellidae
493
20a(19b)
Dorsum with white genital and anal patches, one or more medial white patches and a white bar on neck; no papillae on dorsum;
five longitudinal rows of white prominences surrounded by yellowish dots, equidistant longitudinally and transversely; body very
flattened; length 10 mm .....................................................................................................Desserobdella michiganensis Sawyer, 1972
20b.
Dorsum without white patches and bar on neck ..............................................................................................................................21
21a(20b)
Posterior sucker separated from body on a short pedicel; dorsum smooth; eight rows of inconspicuous sensillae on dorsal annuli;
only recorded parasitism on Cryptobranchus alleganiensis; length 17 mm .....................................................................Batracobdella
(Desserobdella) cryptobranchii Johnson and Klemm, 1977
21b.
Posterior sucker not on pedicel; posterior sucker small; low small papillae present; 3-annulate (Fig. 3a); length 25 mm ..................22
22a(21b)
Dorsal papillae present; lateral metameric and dorsal color markings; 60 radiating papillaeon rim of posterior sucker; feeds mainly
on fish; length 10 mm .....................................................................................................................Deserobdella phalera (Graf, 1899)
22b.
No dorsal papillae; no lateral metameric and dorsal colour markings; no papillae on rim of posterior sucker; feeds mainly on amphibians; length 5 mm ....................................................................................................................Desserobdella picta (Verrill, 1872)
23a(19b)
Brown, horny, chitinous scute (nuchal plate) on the dorsal surface of segment VIII (Fig. 24) ...........................................................24
23b.
Without a nuchal plate .....................................................................................................................................................................25
24a(23a)
Pigment pattern arranged in longitudinal stripes on dorsum; six branched pairs of crop ceca; posterior sucker pigmented on dorsum; length 18 mm ...................................................................................................................Helobdella california Kutschera, 1988
24b.
No longitudinal stripes on dorsum; six unbranched pairs of crop ceca, the last pair posteriorly; posterior sucker unpigmented;
length 14 mm ............................................................................................................................Helobdella stagnalis (Linnaeus, 1758)
25a(23b)
Dorsal surface smooth......................................................................................................................................................................26
25b.
Dorsal surface with 3 – 7 longitudinal series of papillae, or with scattered papillae...........................................................................28
26a(25a)
Body unpigmented and translucent; body rounded and subcylindrical; lateral margins almost parallel; posterior sucker small and
terminal; one pair of crop ceca; length 25 mm ...............................................................................Helobdella elongata (Castle, 1900)
26b.
Body pigmented, with or without longitudinal or transverse bands; body flat with posterior wider than tapering anterior; six pairs
of crop ceca ......................................................................................................................................................................................27
27a(26b)
Dorsum without transverse pigmentation; dorsum with six prominent longitudinal white stripes alternating with brown stripes;
length 14 mm ......................................................................................................................................Helobdella fusca (Castle, 1900)
27b.
Dorsum with transverse brown interrupted stripes alternating with irregular white banks; length 10 mm ...........................................
Helobdella transversa Sawyer, 1972
28a(25b)
Dorsal surface with 5 – 9 longitudinal rows of papillae; papillae large, conspicuous, and rounded; lightly or unpigmented; length 14
mm ...............................................................................................................................................Helobdella papillata (Moore, 1906)
28b.
Dorsal surface with three or fewer incomplete series of small papillae or with scattered papillae; length 25 – 30 mm .......................29
29a(28b)
Ratio of body width to body length 0.26 0.02 SD. Dorsal surface with longitudinal strips many of which are interrupted by
circular zones of unpigmented skin. Five pairs of crop caeca which have numerous secondary diverticula with a bumpy
outline ....................................................................................................................................Helobdella triserialis (Blanchard, 1849)
29b.
Ratio of body width to body length 0.37 0.01 SD. Dorsal surface with five narrow longitudinal stripes which extend unbroken
along the body. Five pairs of crop ceca which are bilobed and smooth in outline ...................................................Helobdella robusta
Shankland, Martindale, Nardelli-Haefinger, Baxter and Price, 1992
30a(6b)
Four pairs of eyes on paramedian lines of segments II – V (Fig. 25a); body very soft.....................................Theromyzon................31
30b.
Three pairs of eyes (Fig. 25b, c) (coalesced eyes sometimes occur, but the lobed nature indicates the original condition); body
firm ..................................................................................................................................................................................................35
31a(30a)
Gonopores separated by two annuli .................................................................................................................................................33
31b.
Gonopores separated by three or four annuli ...................................................................................................................................32
32a(31b)
Gonopores separated by three annuli cocoons attached to ventral body wall, two female pores.......................................Theromyzon
trizonare Davies and Oosthuizen, 1992
32b.
Gonopores separated by four annuli cocoons attached directly to substrate, single female pore .......................................Theromyzon
tessulatum (Müller, 1776)
33a(31a)
Two female pores ..................................................................................................................Theromyzon maculosum (Rathke, 1862)
33b.
One female pore ...............................................................................................................................................................................34
494
Ronald W. Davies and Fredric R. Govedich
34a(33b)
Female atrium cylindrical ....................................................................................Theromyzon bifarium Oosthuizen and Davies, 1993
34b.
Female atrium spherical ......................................................................................................................Theromyzon rude (Baird, 1869)
35a(30b)
First pair of eyes closer together than succeeding two pairs, i.e., eyes arranged in triangular pattern (Fig. 25c); no papillae; male
and female ducts open into a common gonopore; little pigmentation; generally amber colored; length 10 mm ....................................
Alboglossiphonia heteroclita (Linnaeus, 1761)
35b.
Eyes equidistant in two paramedian rows (Fig. 25b) ........................................................................................................................36
36a(35b)
Dorsum sometimes with papillae on annulus a2 in six longitudinal rows; pair of paramedial stripes on dorsum and ventrum; six
pairs of crop ceca; gonopores separated by two annuli (Fig. 3b); length 25 mm................Glossiphonia complanata (Linnaeus, 1758)
36b.
Dorsum with large, distinct papillae on annuli a2 and a3; dorsum with numerous, irregularly shaped whitish spots; seven pairs of
crop ceca; length 25 mm ............................................................................................................Boreobdella verrucata (Müller, 1844)
37a(2b)
Posterior sucker flattened, as wide or wider than the widest part of body (Fig. 1b); pulsatile vesicles on lateral margins of neural annuli of urosome (Fig. 6a, b); zero or two pairs of eyes ......................................................................................................................38
37b.
Posterior sucker concave, weakly developed, and narrower than widest part of the body; no pulsatile vesicles (Fig. 26); zero or one
pair of eyes .......................................................................................................................................................................................45
38a(37a)
Body divided into anterior trachelosome and posterior urosome; 11 pairs of pulsatile vesicles not very apparent in preserved specimens; each pulsatile vesicle covers two annuli; body cylindrical or sometimes slightly flattened; two pairs of eyes; both anterior and
posterior suckers wider than body; oculiform spots present on posterior sucker (Fig. 19b, c, d) )(except Piscicola punctata) (Fig.
19a): 14-annulate (except Piscicola punctata which is 3-annulate) ...............................................................Piscicola......................39
38b.
Body divided into distinct anterior trachelosome and posterior urosome; 11 pairs of large and distinct pulsatile vesicles easily seen
in preserved and live specimens (Figs. 1b and 6); each pulsatile vesicle covers four annuli; well-developed anterior and posterior
suckers; no papillae; zero or two pairs of eyes (Fig. 6); oculiform spots on posterior sucker present or absent (Fig. 9a); 7-annulate;
length 80 mm .......................................................................................................................Cystobranchus.....................................42
39a(38a)
Posterior sucker with 8 – 14 oculiform spots (Fig. 19b, c, d) .............................................................................................................40
39b.
Posterior sucker without oculiform spots (Fig. 19a); 3-annulate (Fig. 2a); two pairs of crescent-shaped eyes: gonopores separated by
three or four annuli; length 16 mm ...................................................................................................Piscicola punctata (Verrill, 1872)
40a(39a)
Posterior sucker with 10 – 14 punctiform oculiform spots (Fig. 19c, d).............................................................................................41
40b.
Eight to ten crescent-shaped oculiform spots on the posterior sucker (Fig. 19b); gonopores separated by two annuli; 14-annulate
length 31 mm...................................................................................................................................Piscicola salmositica Meyer, 1946
41a(40a)
With 10 – 12 (usually 10) punctiform oculiform spots on posterior sucker; dark rays absent from posterior sucker (Fig. 19c): gonopores separated by two annuli (Fig. 3a); anterior pair of eyes like heavy dashes twice as long as wide; length 24 mm Piscicola milneri
(Verrill, 1874)
41b.
With 12 – 14 punctiform, oculiform spots on posterior sucker, separated by an equal number of dark pigmented rays (Fig. 19d);
gonopores separated by three annuli; anterior pair of eyes like fine dashes five times as long as wide; length 30 mm ..Piscicola geometra (Linnaeus, 1758)
42a(38b)
With oculiform spots on posterior sucker.........................................................................................................................................43
42b.
Without oculiform spots on posterior sucker....................................................................................................................................44
43a(42a)
Eight oculiform spots on posterior sucker; two rows of 12 lateral oculiform spots on each side of body (Fig. 6b); two pairs of eyes;
length 7 mm ..............................................................................................................Cystobranchus meyeri Hayunga and Grey, 1976
43b.
Ten oculiform spots on posterior sucker; no lateral ocelli; two pairs of eyes; length 15 mm..........................Cystobranchus virginicus
Hoffman, 1964
44a(42b)
Two pairs of eyes; the first pair forming conspicuous dashes at 45° to the longitudinal axis; the second part of eyes ovoid (Fig. 6a);
gonopores separated by two annuli (Fig. 3b); length 30 m ............................................................Cystobranchus verrilli Meyer, 1940
44b.
No eyes; length 30 mm......................................................................................................Cystobranchus mammillatus (Malm, 1863)
45a(37b)
Body divided into small trachelosome and larger urosome (Fig. 26b); length-to-width ratio 3 – 6:1; one pair of eyes; 12- or 14-annulate ....................................................................................................................................................Myzobdella lugubris Leidy, 1851
45b.
Body not divided into trachelosome and urosome (Fig. 26a); zero or one pair of eyes......................................................................46
46a(45b)
Length-to-width ratio 10:1; zero or one pair of eyes in posterior half of the head; 12- or 14-annulate.......................Illinobdella 47
46b.
Length-to-width ratio 4 – 5:1 (Fig. 26b); one pair of eyes; 3-annulate (Fig. 2a); dorsum with six black / brown longitudinal stripes;
length 8 mm ......................................................................................................................................Piscicolaria reducta Meyer, 1940
13. Annelida: Euhirudinea and Acanthobdellidae
495
47a(46a)
Length-to-width ratio about 10:1; 14-annulate ......................................................................................Illinobdella alba Meyer, 1940
47b.
Length-to-width ratio 15:1 ............................................................................................................................................................48
48a(47b)
Length-to-width ratio 15:1; one pair of eyes (sometimes absent); anus 15 annuli anterior to posterior sucker; 14-annulate; posterior
sucker little more than concavity of the posterior body ...............................................................Illinobdella richardsoni Meyer, 1940
48b.
Length-to-width-ratio 18:1; one pair of eyes; anus ten annuli or less anterior to posterior sucker; 12-annulate; posterior sucker half
as wide as posterior body ................................................................................................................Illinobdella elongata Meyer, 1940
49a(3a)
With copulatory gland pores on the ventral surface, 10 or 11 annuli posterior to the male gonopores (Figs. 27 and 28) .....................
Macrobdella .....................................................................................................................................................................................50
49b
Ventral copulatory gland pairs absent ..............................................................................................................................................53
50a(49a)
With 24 copulatory gland pores (four rows with six gland pores each) (Fig. 28a) on raised pads; dorsum with red / orange spots;
two to two-and-a-half annuli between gonopores; length 150 mm ............................................Macrobdella sestertia Whitman, 1886
50b.
With four, six, or eight copulatory gland pores (Fig. 28); with or without red / orange spots on dorsum ..........................................51
51a(50b)
Eight copulatory gland pores (Fig. 28b) (2 rows of 4); two annuli between gonopores; dorsum without red / orange spots; length
150 mm ...........................................................................................................................................Macrobdella ditetra Moore, 1953
51b.
Four or six copulatory gland pores; dorsum with red / orange spots .................................................................................................52
52a(51b)
Four copulatory gland pores (Fig. 27) (two rows of two); five to five and-a-half annuli between gonopores; jaws well developed
with about 65 monostichodont acute teeth (Fig. 9a); 10 pairs of testisacs; ventrum red / orange; length 150 mm .................................
Macrobdella decora (Say, 1824)
52b.
Six copulatory gland pores (Fig. 28c) (three rows of two); four or five annuli between gonopores; ventrum yellow / grey; length
150 mm ......................................................................................................................................Macrobdella diplotertia Meyer, 1975
53a(49b)
Glandular area around gonopores; gonopores separated by three or four annuli .................................................................................
Philobdella .......................................................................................................................................................................................54
53b.
No glandular area around gonopores; gonopores separated by 5 – 7 annuli......................................................................................55
54a(53a)
20 – 26 distichodont teeth per jaw (Fig. 9b); lateral margins of dorsum without discrete spots; length 85 mm....................Philobdella
floridana (Verrill, 1874)
54b.
35 – 48 distichodont teeth per jaw; seven pairs of testisacs; irregular black spots on margins of dorsum; length 85 mm.......................
Philobdella gracilis Moore, 1901
55a(53b)
Jaws absent (Fig. 8a) ........................................................................................................................................................................56
55b.
Jaws present (Fig. 8b, c) and denticulate (Fig. 9) ..............................................................................................................................57
FIGURE 27 Ventral view of the male and female gonopores of Macrobdella decora. FIGURE 28 Diagrammatic representation of the relative slopes and positions of the male and female gonopores and copulatory
glands of: (a) Macrobdella sestertia; (b) Macrobdella ditetra; and (c) Macrobdella diplotertia.
496
Ronald W. Davies and Fredric R. Govedich
56a(55a)
Lower surface of velum smooth (Fig. 8a); gonopores in the furrows between the annuli and separated by five annuli; pharynx with
12 internal ridges; length 300 mm...................................................................................................Mollibdella grandis (Verrill, 1874)
56b.
Lower surface of velum papillate; gonopores in the middle of annuli and separated by five annuli; pharynx with 15 internal ridges
................................................................................................................................................Bdellarogatis plumbeus (Moore, 1912)
57a(55b)
Jaws small and retractable into narrow-mouthed tubular pits; 9 – 25 distichodont (Fig. 9b) teeth per jaw............................................
Percymoorensis ................................................................................................................................................................................58
57b.
Jaws large; 35 – 100 acute monostichodont (Fig. 9a) teeth; length 100 mm ...................................Hirudo medicinalis Linnaeus, 1758
58a(57a)
Gonopores separated by five to five-and-a-half annuli; female gonopore small.................................................................................59
58b.
Gonopores separated by six and-a-half to seven annuli; female gonopore large and conical length 200 mm ........................................
Percymoorensis septagon Sawyer and Shelley, 1976
59a(58a)
Dorsum with median black stripe.....................................................................................................................................................60
59b.
Dorsum with irregular, scattered black spots ....................................................................................................................................61
60a(59a)
Dorsum uniformly black or slate gray; posterior sucker narrower than body; jaws with 20 – 25 pairs of teeth; length 250 mm ...........
Percymoorensis lateralis (Say, 1824)
60b.
Dorsum brown / green; posterior sucker as wide as body; 9 – 14 pairs teeth ................................Percymoorensis kingi (Mather, 1954)
61a(59b)
Jaws with 10 – 12 teeth; posterior sucker about 75% of maximal body width and attached by very short pedicel; length 75 mm ........
...................................................................................................................................Percymoorensis lateromaculata (Mather, 1963)
61b.
Jaws with 12 – 16 pairs of teeth; posterior sucker about half maximum body width; length 150 mm...........................Percymoorensis
marmorata (Say, 1824)
62a(3b)
Somites 5-annulate (b1, b2, a2, b5, b6) (Fig. 29a) with all annuli equal in length; three pairs of eyes; gonopores separated by two annuli (Fig. 3b); length 100 mm ...........................................................................................................................................................63
62b.
Somites six or seven-annulate (Fig. 29b); annuli of unequal length units will be either subdivided or longer than the others; in any
group of six consecutive annuli at least one annulus narrower or wider than the others; three or four pairs of eyes ........................65
63a(62a)
Eyes all similar in size; annuli not raised on dorsum and without papillae; anus located three or four segments anterior to posterior
sucker; mouth small; gonopores in furrows separated by 2 – 5 annuli .............................................Erpobdella punctata (Leidy, 1870)
63b.
Eyes differ in size with the second and third pairs smaller than the anterior pair; each annulus raised on dorsum with 10 – 18 small
white-tipped papillae; anus located at base of posterior sucker; one or two pairs of crop ceca gonopores in furrows separate by two
annuli...............................................................................................................................................Motobdella..............................64
64a(63b)
14 – 18 small, white-tipped papillae in a ring about each annulus; mouth very large (4.0 mm); two pairs of crop ceca nephridiopores
not visible, internal surface of posterior sucker pigmented ...........................Motobdella montezuma Davies, Singhal and Blinn, 1985
FIGURE 29
(a) Annulation of Erpobdella with all annuli (b1, b2, a2, b5, b6) equal in length; and
(b) Nephelopsis, Dina, or Mooreobdella with the annuli (b1, b2, a2, b5, c11, c12) of unequal lengths.
13. Annelida: Euhirudinea and Acanthobdellidae
497
64b.
10 – 14 small, white-tipped papillae on dorsal surface only of each annulus; mouth 1.1 mm in diameter; nephridiopores visible as
small white spots; internal surface of posterior sucker not pigmented; one or two pars of crop ceca.................Motobdella sedonensis
Govedich, Blinn, Keim and Davies, 1998
65a(62b)
Four pairs of eyes, two labial and two buccal of similar size ............................................................................................................66
65b.
Zero or three pairs of eyes ....................................................................................................................................Dina / Mooreobdella
.........................................................................................................................................................................................................68
66a(65a)
Two annuli between gonopores (Fig. 3a); anus one segment anterior to poster sucker; atrial cornua spirally coiled like the horn of a
ram (Fig. 18a); length 100 mm ........................................................................................................Nephelopsis obscura Verrill, 1872
66b.
Three or more annuli between gonopores; atrial cornua simply curved (Fig.18b) .........................................................................Dina
.........................................................................................................................................................................................................67
67a(66b)
Three-and-a-half to four annuli between gonopores; body heavily blotched with a median stripe; anus large, opening on a conical
tubercle; length 60 mm..............................................................................................................Dina dubia (Moore and Meyer, 1951)
67b.
Three to three-and-a-half annuli between gonopores; nearly pigmentless or with a few dark spots; anus small, not on tubercle;
length 30 mm ................................................................................................................................................Dina parva Moore, 1912
68a(65b)
Dorsum lacking scattered black pigment ..........................................................................................................................................69
68b.
Dorsum with scattered black pigment; two annuli between gonopores (Fig. 3a); length 55 mm .....................................Mooreobdella
melanostoma Sawyer and Shelley, 1976
69a(68a)
Two to two-and-a-half annuli between gonopores............................................................................................................................70
69b.
Three to four-and-a-half annuli between gonopores .........................................................................................................................72
70a(69a)
Male gonopore surrounded by circle of papillae; zero or three pairs of eyes; two annuli between gonopores (Fig. 3a); length 15 mm
Dina anoculata Moore, 1898
70b.
No papillae around male gonopore; three (rarely four) pairs of eyes ................................................................................................71
71a(70b)
Two annuli between gonopores (Fig. 3a); three (rarely four) pairs of eyes; length 50mm................................................Mooreobdella
fervida Verrill, 1872
71b.
Either 2 or 2 1/2 annuli between gonopores; three pairs of eyes; length 30 mm.............................Mooreobdella bucera Moore, 1949
72a(69b)
Three pairs of eyes; gonopores separated by three annuli; length 50 mm ............................Mooreobdella microstoma (Moore, 1901)
72b.
Gonopores separated by 4 – 4 1/2 annuli; three pairs of eyes; length 40 mm.............Mooreobdella tetragon Sawyer and Shelley, 1976
ACKNOWLEDGMENTS
It is a pleasure to acknowledge the technical assistance and patience of Mrs Thelma Thompson in the preparation of this chapter.
We would also like to acknowledge Dr. Bonnie Bain for her assistance in proofreading and reviewing this chapter and key.
VIII. LITERATURE CITED
Able, K. W. 1976. Cleaning behaviour in the cyprinodont fishes: Fundus majalis, Cyprinodon variegatus and Lucania parva. Chesapeake Science 17:35 – 39.
Anderson, R. S., Raasveldt. L.G 1974. Gammarus predation and the
possible effects of Gammarus and Chaoborus feeding on the
zooplankton composition in some small lakes and ponds in
western Canada. Canadian Wildlife Service Occasional Paper
No. 18.
Angstadt, J. D., Moore. W.H. 1997. A circadian rhythm of swimming behavior in a predatory leech of the Family Erpobdellidae.
The American Midland Naturalist 137:165 – 172.
Anholt, B. R. 1986. Prey selection by the predatory leech Nephelopsis obscura in relation to three alternative models of foraging.
Canadian Journal of Zoology 64:649 – 555.
Anholt, B. R., Davies, R. W. 1987. Effects of hunger level on the activity of the predatory leech Nephelopsis obscura Verrill
(Hirudinoidea: Erpobdellidae). American Midland Naturalist
177:307 – 311.
Arnold, S. J. 1981. Behavioural variation in natural populations. 1.
Phenotypic, genetic and environmental correlations between
chemoreceptive responses to prey in the garter snake
Thamnophis elegans. Evolution 35:489 – 509.
Babin, J., Prepas. E. E. 1985. Modeling winter oxygen depletion
rates in ice-covered temperate zone lakes in Canada. Canadian
Journal of Fisheries and Aquatic Sciences 42:239 – 249.
Baird, D. J., Linton, L. R., Davies. R. W. 1986. Life-history evolution
and post-reproductive mortality risk. Journal of Animal Ecology 55:295 – 302.
Baird, D. J., Gates, T. E., Davies. R. W. 1987a. Oxygen conditions in
two prairie pothole lakes during winter ice cover. Canadian
Journal of Fisheries and Aquatic Sciences 44:1092 – 1095.
Baird, D. J., Linton, L. R., Davies. R. W. 1987b. Life-history flexibility as a strategy for survival in a variable environment. Functional Ecology 1:45 – 48.
Baird, W. 1869. Descriptions of some new suctorial annelids in the
collection of the British Museum. Proceedings of the Zoological
Society of London. pp. 310 – 318.
Barta, J. R., Sawyer, R. T. 1990. Definition of a new genus of
glossiphoniid leech and a redescription of the type species,
498
Ronald W. Davies and Fredric R. Govedich
Clepsine picta Verrill, 1872. Canadian Journal of Zoology
68:1942 – 1950.
Bartonek, J. C. 1972. Summer foods of American Widgeon, Mallards
and Green-winged Teal near Great Slave Lake, Northwest
Territories. Canadian Field Naturalist 86:373 – 376.
Bartonek, J. C., Hickey, J. J. 1969a. Selective feeding by juvenile
diving ducks in summer. The Auk 86:457493.
Bartonek, J. C., Hickey, J. J. 1969b. Food habits of canvasbacks, redheads and lesser scaup in Montana. Condor 71:280 – 290.
Bartonek, J. C., Murdy, H. W. 1970. Summer foods of lesser scaup in
subarctic taiga. Arctic 23:35 – 44.
Bartonek, J. C., Trauger, D. L. 1975. Leeches (Hirudinea) infestations
among waterfowl near Yellowknife, Northwest Territories.
Canadian Field Naturalist 89:234 – 243.
Beadle, L. D. 1939. Regulation of the haemolymph in the saline water mosquito larva Aedes detritus (Edw.). Journal of Experimental Biology 16:346 – 362.
Bendell, B. E., McNicol, D. K. 1991. An assessment of leeches
(Hirudinea) as indicators of lake acidification. Canadian Journal of Zoology 69:130 – 133.
Bennike, S. A. B. 1943. Contributions to the ecology and biology of
the Danish freshwater leeches. Folia Limnologica Scandinavica
2:1 – 109.
Biernacka, B., Davies, R. W. 1994. The effects of temperature and
feeding regime on hermaphroditism and gametogenesis of
Nephelopsis obscura, a freshwater predatory leech. Canadian
Journal of Zoology 72:1639 – 1642.
Biernacka, B., Davies, R. W. 1995. Is a visible clitellum an index of
sexual maturity in erpobdellid leeches? Invertebrate Reproduction and Development 28:97 – 101.
Blair, W. N. 1927. Notes on Hirudo medicinalis, the medicinal leech,
as a British species. Proceedings of the Zoological Society of
London 11:999 – 1002.
Blanchard, E. 1849. Annelides. Hirudineanos. Gay’s historia fisca y
politico de Chile. Zoologia, Paris 3:43 – 50.
Blanchard, R. 1893. Courtes notices sur les Hirudinees: X. Hirudinees de l’Europe boreale. Bulletin Societe Zoologique de France
18:93 – 94.
Blinn, D. W., Davies, R. W. 1989. The evolutionary importance of
mechanoreception in three erpobdellid leech species. Oecologia
79:6 – 9.
Blinn, D. W., Davies, R. W. 1990. Concomitant diel vertical migration of a predatory leech and its amphipod prey. Freshwater
Biology 24:401 – 407.
Blinn, D. W., Davies, R. W., Dehdashti, B. 1987. Specialized pelagic
feeding by Erpobdella montezuma (Hirudinea). Holarctic Ecology 10:235 – 340.
Blinn. D. W., Pinney, C. Wagner, V. T. 1988. Intraspecific discrimination of amphipod prey by a freshwater leech through
mechanoreception. Canadian Journal of Zoology 66:427 – 430.
Blinn, D. W., Dehdashti, B. Runck, C. Davies, R. W. 1990. The importance of prey size and density in an endemic predator – prey
couple (leech Erpobdella montezuma – amphipod Hyalella montezuma). Journal of Animal Ecology 59:187 – 192.
Briggs, D. E. 1991. Extraordinary fossils. American Scientist
79:130 – 141.
Bulmer, M. G. 1985. Selection for iteroparity in a variable environment. American Naturalist 136:63 – 71.
Bur, M.T. 1994. Incidence of the leech Actinobdella pediculata on
freshwater drum in Lake Erie. Journal of Great Lakes Research
20:768 – 770.
Büsing, K. H. 1951. Pseudomonas hirudinis, ein bakterieller Darmsybiont des Blutegels (Hirudo offcinalis). Zentralblatt fuer Bakterienologie Parasitenkunde 157:478 – 484.
Büsing, K. H., Döll, W., Freytag, K. 1953. Die Bakterienflora der
medizinischen Blutegel. Archiv fur Mikrobiologie 19:52 – 86.
Calow, P., Riley, H. 1982. Observations on reproductive effort in
British erpobdellid and glossiphoniid leeches with different life
cycles. Journal of Animal Ecology 51:697 – 712.
Carena, H. 1824. Monographic due genre Hirudo. Supplement.
Memoire Accademia Science, Torino 28:331 – 337.
Casselman, J. M., Harvey, H. H. 1975. Selective fish mortality resulting from low winter oxygen. Internationale Vereinigung für
Theoretische und Angewandte Limnologie 19:2418 – 2429.
Castle, W. E. 1900. Some North American fresh-water Rhynchobdellidae, and their parasites. Bulletin of the Museum of Comparative Zoology. Harvard 36:17 – 64.
Croghan, P. C. 1958. The survival of Artemia salina (Linn.) in various media. Journal of Experimental Biology 35:213 – 216.
Cywinska, A., Davies, R. W. 1989. Predation on the erpobdellid
leech Nephelopsis obscura in the laboratory. Canadian Journal
of Zoology 67:2689 – 2693.
Daborn, G. R. 1976. Colonization of isolated aquatic habitats.
Canadian Field-Naturalist 90:56 – 57.
Dahl, J. 1998. The impact of vertebrate and invertebrate predators
on a stream benthic community. Oecologia 117:217 – 226.
Dahl, J., Greenberg, L. 1997. Foraging rates of a vertebrate and an
invertebrate predator in stream enclosures Oikos 78:459 – 466.
Dahm, A. G. 1962. Distribution and biological patterns of Acanthobdella peledina Grube from Sweden (Hirudinea, Acanthobdellidae). Acta Universitatis Lundensis Arrskrift 58:1 – 35.
Daniels, B. A., Sawyer R. T. 1973. Host-parasite relationship of the
fish leech Illinobdella moorei Meyer and the white catfish
lctalurus catus (Linnaeus). Bulletin of the Association of Southeastern Biologists 20:48.
Davies, R. W. 1969. The production of antisera for detecting specific
triclad antigens in the gut contents of predators. Oikos
20:248 – 260.
Davies, R. W. 1971. A key to the freshwater Hirudinoidea of
Canada. Journal of the Fisheries Research Board of Canada
28:543 – 552.
Davies, R. W. 1972. Annotated bibliography to the freshwater
leeches (Hirudinoidea) of Canada. Fisheries Research Board of
Canada., Technical Report No. 306.15 pp.
Davies, R. W. 1973. The geographic distribution of freshwater Hirudinoidea in Canada. Canadian Journal of Zoology 51:531 – 545.
Davies, R. W. 1978. Reproductive strategies shown by freshwater
Hirudinoidea. Internationale Vereinigung für Theoretische und
Angewandte Limnologie 20: 2378 – 9381.
Davies, R. W. 1979. Dispersion of freshwater leeches (Hirudinoidea)
to Anticosti Island, Quebec. Canadian Field-Naturalist 93:
310 – 313.
Davies, R. W. 1984. Sanguivory in leeches and its effects on growth,
survivorship and reproduction of Theromyzon rude. Canadian
Journal of Zoology 62:589 – 593.
Davies, R. W. 1987. All about leeches. Nature (London) 325:585.
Davies, R. W. 1991. Annelida: Leeches, Polychaetes, and Acanthobdellids, in: Thorp, J. H. Covich, A. P. Eds. Ecology and Classification of North American Freshwater Invertebrates. Academic
Press, San Diego.
Davies, R. W., Everett, R. P. 1977. The life history, growth and age
structure of Nephelopsis obscura Verrill, 1872 (Hirudinoidea)
in Alberta. Canadian Journal of Zoology 55:620 – 627.
Davies, R. W., Gates, T. E. 1991a. Intra- and interspecific differences
in the response of two lentic species of leeches to seasonal hyperoxia. Canadian Journal of Fisheries and Aquatic Science
48:1124 – 1127.
Davies, R. W., Gates, T. E. 1991b. The effects of different oxygen
13. Annelida: Euhirudinea and Acanthobdellidae
regimes on the feeding and vertical distribution of Nephelopsis
obscura (Hirudinoidea). Hydrobiologia 211: 51 – 56.
Davies, R. W., Kassera, C. E. 1989. Foraging activity of two specis of
predatory leeches exposed to active and sedentary prey. Oecologia 81:329 – 334.
Davies, R. W., Oosthuizen, J. H. 1992. A new species of duck leech
from North America formerly confused with Theromyzon rude
(Rhynchobdellida: Glossiphoniidae. Canadian Journal of Zoology 71:770 – 775.
Davies, R. W., Reynoldson, T. B. 1976. A comparison of the life-cycle of Helobdella stagnalis (Li. 1758) (Hirudinoidea) in two different geographic areas in Canada. Journal of Animal Ecology
45:457 – 470.
Davies, R. W., Singhal, R. N. 1988. Cosexuality in the leech, Nephelopsis obscura (Erpobdellidae). International Journal of Invertebrate Reproduction and Development 13:55 – 64.
Davies, R. W., Wilkialis, J. 1982. Observations on the ecology and
morphology of Placobdella papillifera (Verrill) (Hirudinoidea:
Glossiphoniidae) in Alberta, Canada. American Midland Naturalist 107:316 – 324.
Davies, R. W., Reynoldson, T. B., Everett, R. P. 1977. Reproductive
strategies of Erpobdella punctata (Hirudinoidea) in two temporary ponds. Oikos 29:313 – 319.
Davies, R. W., Wrona, F. J., Everett, R. P. 1978. A serological study
of prey selection by Helobdella stagnalis (Hirudinoidea). Journal of Animal Ecology 48:181 – 194.
Davies, R. W., Wrona, F. J., Linton, L., Wilkialis, J. 1981. Inter- and
intra-specific analyses of the food niches of two sympatric
species of Erpobdellidae (Hirudinoidea) in Alberta, Canada.
Oikos 37:105 – 111.
Davies, R. W., Linton, L. R., Parsons, W., Edgington, E. S. 1982a.
Chemosensory detection of prey by Nephelopsis obscura
(Hirudinoidea: Erpobdellidae). Hydrobiologia 97:157 – 161.
Davies, R. W., Linton, L. R. Wrona, F. J. 1982b. Passive dispersal of
four species of freshwater leeches (Hirudinoidea) by ducks.
Freshwater Invertebrate Biology 1:40 – 44
Davies, R. W., Wrona, F. J. Linton, L. 1982c. Changes in numerical
dominance and its effects on prey utilization and interspecific
competition between Erpobdella punctata and Nephelopsis obscura (Hirudinoidea): an assessment. Oikos 39:92 – 99.
Davies, R. W., Singhal, R. N. Blinn, D. W. 1985. Erpobdella montezuma (Hirudinoidea: Erpobdellidae), a new species of freshwater leech from North America. Canadian Journal of Zoology
63:965 – 969.
Davies, R. W., Yang, T., Wrona, F. J. 1987. Inter- and intra-specific
differences in the effects of anoxia on erpobdellid leeches using
static and flow-through systems. Holarctic Ecology 10:149 – 153.
Davies, R. W., Blinn, D. W., Dehdashti, B., Singhal, R. N. 1988. The
comparative ecology of three species of Erpobdellidae (Annelida-Hirudinoidea). Archiv fur Hydrobiologie 111:601 – 614.
Davies, R. W., Monita, D. M. A. Dratnal, E. Linton, L. R. 1992. The
effects of different oxygen regimes on the growth of a freshwater leech. Ecography 15:190 – 194.
Davies, R .W., Singhal, R. N., Wicklum D. D. 1995. Changes in
reproductive potential of the leech Nephelopsis obscura (Erpobdellidae) as biomarkers for cadmium stress. Canadian Journal
of Zoology 73:2192 – 2196.
Davies, R. W., Dratnal, E., Linton, L. R. 1996. Activity and foraging
behaviour in the predatory freshwater leech Nephelopsis obscura (Erpobdellidae). Functional Ecology 10:51 – 54.
De Eguileor, M., Daniel, S., Giordana, B., Lanzavecchia, G., Valvassori, R. 1994. Trophic exchanges between parent and young
during development of Glossiphonia complanata (Annelida,
Hirudinea). Journal of Experimental Zoology 269:389 – 402.
499
De Filippi, F. 1849. Sopra un nuovo genere (Hlaementeria) di Annelidi della famigliadelle Sanguisughe. Memoires de Academie
de Science., Tonno 10:1 – 14.
Derosa, X. S., Friesen, W. O. 1981. Morphology of leech sensilla: observations with the scanning electron microscope. Biological
Bulletin 160:383 – 393.
Dickinson, M. H., Lent, C. M. 1984. Feeding behaviour of the
medicinal leech, Hirudo medicinalis L. Journal of Comparative
Physiology 154:449 – 455.
Dieter, C. D., Duffy, W. G., Flake, L. D. 1996. The effect of phorate
on wetland macroinvertebrates. Environmental Toxicology and
Chemistry 15:308 – 312.
Dratnal, E., Davies, R. W. 1990. Feeding and energy allocation in
Nephelopsis obscura following exposure to winter stress. Journal of Animal Ecology 59:763 – 773.
Dratnal, E., Dratnal, P. A., Davies, R. W. 1992. The effect of food
availability and foraging constraints on the life history of a
predatory leech, Nephelopsis obscura. Journal of Animal Ecology 61:373 – 379.
Dratnal, E., Reddy, D. C., Biernacka, B., Davies, R. W. 1993. Facultative physiological adaptation and compensation to winter
stresses in the predatory leech Nephelopsis obscura. Functional
Ecology 7:91 – 96.
Ekman, S. 1911. Die Bodenfauna des Vattern, Qualitative und Quantitativ Untersucht. Internationale Revue der gesamten Hydrobiologie 7:146 – 204.
Elliott, J. M. 1973. The diel activity pattern, drifting and food
of the leech Erpobdella octoculata (L.) (Hirudinea: Erpobdellidae) in a Lake District stream. Journal of Animal Ecology
42:449 – 459.
Elliott, J. M., Mann, K. H. 1979. A key to the British freshwater
leeches. Freshwater Biological Association Scientific Publication
No. 40,72 pp.
Fauvel, P. 1922. Un nouveau serpulien d’eau Saumatre Mercierella
enigmatica. Bulletinde la Societe Zoologique de France 47:
425 – 431.
Fernandez, J. 1980. Embryonic development of the Glossiphoniid
leech Theromyzon rude: characterization of developmental
stages. Developmental Biology 76:245 – 262.
Fernandez, J., Tellez, V., Olea, N. 1992. Hirudinea. Microscopic
Anatomy of Invertebrates 7:323 – 394.
Frey, H., Leuckart, R. 1847. Verzeichnis der zur Fauna Helgolands
gehörenden wirbellosen seethiere. Beiträgen zur Kenntniss
wirbelloser Thiere, pp. 136 – 168.
Friesen, W. O. 1981. Physiology of water motion detection in the
medicinal leech. Journal of Experimental Biology 92:255 – 275.
Friesen, W. O., Dedwylder, R. D. 1978. Detection of low-amplitude
water movements: a new sensory modality in the medicinal
leech. Neuroscience Abstracts 4:380.
Gates, T. E., Davies, R. W. 1987. The influence of temperature on the
depth distribution of sympatric Erpobdellidae (Hirudinoidea).
Canadian Journal of Zoology 65:1243 – 1246.
Gates, T. E., Baird, D. J., Wrona, F. J., Davies, R. W. 1987. A device
for sampling macroinvertebrates in weedy ponds. Journal of the
North American Benthological Society 6:133 – 139.
Gee, W. 1913. The behavior of leeches with special reference to its
modifiability. University of California Publications in Zoology
11:197 – 305.
Gerking, S. D. 1957. A method of sampling the littoral macrofauna
and its application. Ecology 38:219 – 266.
Giesel, J. T. 1974. The biology and adaptability of natural populations. Mosley, St. Louis, Mio.
Govedich, F. R., Blinn, D. W. Hevley, R. H., Keim, P. S. 1999.
Cryptic radiation in erpobdellid leeches in xevic landscapes: a
500
Ronald W. Davies and Fredric R. Govedich
molecular analysis of population differentiation. Canadian
Journal of Zoology 77:52 – 57.
Govedich, F. R., Blinn, D. W., Keim, P., Davies, R. W. 1998. Phylogenetic relationships of three genera of Erpobdellidae (Hirudinoidea), with a description of a new genus, Motobdella, and
species, Motobdella sedonensis. Canadian Journal of Zoology
76:2164 – 2171.
Graf, A. 1899. Hirudineenstudien. Acta Acadamemiae Leopoldensia,
Halle 72:215 – 404.
Grantham, B. A., Hann, B. J. 1994. Leeches (Annelida: Hirudinea) in
the Experimental Lakes Area, Northwestern Ontario, Canada:
Patterns of species composition in relation to environment.
Canadian Journal of Fisheries and Aquatic Science
51:1600 – 1607.
Green, R. H. 1979. Sampling design and statistical methods for environmental biologists. Wiley, New York.
Grube, A. E. 1851. Annulaten reise in den Aussersten Norden und
Ostens Sibiriens. Herausgegeben von Dr. A. Th. Middendorff,
St. Petersburg 1:1 – 254.
Hagadorn, I. R. 1969. Hormonal control of spermatogenesis in
Hirudo medicinalis. II. Testicular response to brain removal
during the phase of testicular maturity. Genetics and Comparitive Endocrolology 12:469 – 478.
Hartman, O. 1951. The literature of the polychaetous annelids. Vol
1. Allan Hancock Foundation. University of Southern California, Los Angeles, 290 pp.
Hartman, O. 1952. Iphitime and Ceratocephala (polychaetous annelids) from California. Bulletin of the Southern California
Academy of Science 51:9 – 20.
Hayunga, E. G., Grey, A. J. 1976. Cystobranchus meyeri sp. n.
(Hirudinea: Piscicolidae) from Catostomus commersoni Lacepede in North America. Journal of Parasitology 62:621 – 627.
Hemingway, E. E. 1908. The anatomy of Placobdella pediculata in:
The leeches of Minnesota. Geological and Natural History of
Minnesota, Zoology Series No. 5, pp. 29 – 63.
Herrmann, S. J. 1970a. Systematics, distribution and ecology of Colorado Hirudinea. American Midland Naturalist 83:1 – 37.
Herrmann, S. J. 1970b. Total residual tolerances of Colorado
Hirudinea. Southwestern Naturalist 15:261 – 273.
Herter, K. 1929a. Temperaturversuche mit Egeln. Zeitschrift fuer
Vergleichende Physiologie 10:248 – 271.
Herter, K. 1929b. Reizphysiologisches Verhalten und Parasitismus
des Entenegels Protoclepsis tesselata O. F. Müller. Zeitschrift
fuer Vergleichende Physiologie10:272 – 308.
Hobbs, H. H., Figueroa, H. V. 1958. The exoskeleton of a freshwater
crab as a microhabitat for several invertebrates. Virginia Journal of Science 9:395 – 396.
Hoffman, R. L. 1964. A new species of Cystobranchus from southwestern Virginia (Hirudinea: Piscicolidae). American Midland
Naturalist 72:390 – 395.
Holmquist, C. 1974. A fish leech of the genus Acanthobdella found
in North America. Hydrobiologia 44:241 – 245.
Holt, P. C. 1969. The systematic position of the Branchiobdellidae
(Annelida: Clitellata). Systematic Zoology 14:25 – 32.
Jennings, J. B., Vanderlande, V. M. 1967. Histochemical and bacteriological studies on digestion in nine species of leeches (Annelida: Hirudinea). Biological Bulletin 133:166 – 183.
Johnson, G. M., Klemm, D. J. 1977. A new species of leech, Batracobdella cryptobranchii n. sp. (Annelida: Hirudinea) parasitic
on the Ozark hillbender. Transactions of the American Microscopical Society 90:331 – 377
Johnson, H. P. 1903. Freshwater nereids from the Pacific coast and
Hawaii, with remarks on freshwater Polychaeta in general.
Mark Anniversary Volume. Holt. New York, pp. 205 – 223.
Jones, S. R .M., Woo, P. T. K. 1990. Redescription of the leech
Desserobdella phalera (Graf, 1899) n. comb. (Rhynchobdellida:
Glossiphoniidae), with notes on its biology and occurrence in
fishes. Canadian Journal of Zoology 68:1951 – 1955.
Kalarani, V., Reddy, D. C., Habibi, H. R., El-Shimy, N., Davies,
R. W. 1995. Occurrence and hormonal action of ecdysone on
gametogenesis and energy utilization in the leech Nephelopsis
obscura (Erpobdellidae). The Journal of Experimental Zoology
273:511 – 518.
Kephart D. G. 1982. Microgeographic variation in the diets of garter
snakes. Oecologia (Berlin) 52:287 – 291.
Kephart, D. G., Arnold, S. J. 1982. Garter snake diets in a fluctuating environment: a seven-year study. Ecology 63:1232 – 1236.
Khan, R. N., Frick, M. G. 1997. Erpobdella punctata (Hirudinea: Erpobdellidae) as phoronts on Ambystoma maculatum (Amphibia:
Ambystomatidae). Journal of Natural History 31: 157 – 161.
Klemm, D. J. 1972a. Freshwater Leeches (Annelida: Hirudinea) of
North America. United States Environmental Protection Agency
Identification Manual No. 8,53 pp.
Klemm, D. J. 1972b. Freshwater polychaetes (Annelida) of North
America. United States Environmental Protection Agency Identification Manual No. 4, 15 pp.
Klemm, D. J. 1977. A review of the leeches (Annelida: Hirudinea) in
the Great Lakes region. Michigan Academician 9:397 – 418.
Klemm, D. J. 1982. Leeches (Annelida: Hirudinea) of North America. United States Environmental Protection Agency
600 / 3 – 82 – 025, 177, pp.
Klemm, D. J. 1985. A Guide to the Freshwater Annelida (Polychaeta,
Naidid and Tubificid Oligochaeta, and Hirudinea) of North
America. Kendall / Hunt, Dubuque, IA. 198 pp.
Krezel, A. M., Wagner, G., Seymour-Ulmer, J., Lazarus, R. A. 1994.
Structure of the RGD protein Decorsin: conserved motif and
distinct function in leech proteins that affect blood clotting. Science 264:1944 – 1947.
Kulkarni, G. K., Nagabhushanen, R., Hanumarte, M. M. 1980. Impact of some neurohumors on the brain neurosecretory profile
of the Indian leech, Poecilobdella viridis (Blanchard). Biological
Journal 2:37 – 41.
Kutschera, U. 1988. A new leech species from North America,
Helobdella californica nov. sp. (Hirudinea: Glossiphoniidae).
Zoologischer Anzeiger 220:173 – 178.
Lasserre, P. 1975. Clitellata, in: Giese A. C. Pearse, J. S. Eds. Reproduction of marine invertebrates. Vol. 3; Annelids: Echiurans.
Academic Press, New York, pp. 215 – 275.
Leidy, J. 1851. Description of Myzobdella. Proceedings of the Academy of Natural Sciences, Philadelphia 1851 – 1853:243.
Leidy, J. 1858. Manavankia speciosa. Proceedings of the Academy of
Natural Sciences, Philadelphia 10:90.
Leidy, J. 1870. Description of Nephelis punctata. Proceedings of the
Academy of Natural Sciences, Philadelphia 22:89.
Light, J. E., Siddall, M. E. 1999. Phylogeny of the leech family Glossiphoniidae based on mitochondrial gene sequences and morphological data. Parasitology 85 (in press).
Linnaeus, C. 1758. Systema naturae. Lipsiae, 10th ed.
Linnaeus, C. 1761. Fauna suecia. Stockholm. 578 pp.
Linton, L. R., Davies, R. W. 1987. An energetics model of an aquatic
predator and its application to life-history optima. Oecologia
71:552 – 559.
Linton, L. R., Davies, R. W., Wrona, F. J. 1982. Osmotic and respirometric responses of two species of Hirudinoidea to changes in
water chemistry. Comparative Biochemistry and Physiology
71A:243 – 247.
Linton, L. R., Davies, R. W., Wrona, F. J. 1983a. The effects of water
temperature, ionic content and total dissolved solids on
13. Annelida: Euhirudinea and Acanthobdellidae
Nephelopsis obscura and Erpobdella punctata (Hirudinoidea,
Erpobdellidae). I. Mortality. Holarctic Ecology 6:59 – 63.
Linton, L. R., Davies, R. W., Wrona, F. J. 1983b. The effect of
water temperature, ionic content and total dissolved solids on
Nephelopsis obscura and Erpobdella punctata (Hirudinoidea,
Erpobdellidae). II. Reproduction. Holarctic Ecology 6:64 – 68.
Lukin, E. I. 1976. Leeches of fresh and brackish water bodies. Fauna
of the Union of Soviet Socialist Republics. Leeches. Vol. 1.
Academy of Science, USSR, Moscow. 484 pp.
Madanmohanrao, G. 1960. Salinity tolerances and oxygen consumption of the cattle leech, Hirudinaria granulosa. Proceedings of
the Indian Academy of Science 11:211 – 218.
Malm, C. W. 1863. Svenska Iglar, Disciferae. Gotheborg. Vetenskapsakademien Arhandlingar i Naturskosarendon 8:153 – 263.
Mann, K. H. 1955. Some factors influencing the distribution of freshwater leeches in Britain. Internationale Vereinigung für Theoretische und Angewandte Limnologie 12:582 – 587.
Mann, K. H. 1956. A study of the oxygen consumption of five
species of leech. Journal of Experimental Biology 33:615 – 626.
Mann, K. H. 1961. Leeches (Hirudinea) their structure, physiology,
ecology and embryology. Pergamon Oxford, 201 pp.
Mannsfeld, W. 1934, in: Literature uber Hirudineen bis zum Jahre
1938. Bronn’s Klassen und Ordnungen des Tierreichs. Bd. 4,
Abt. III: Buch 4. T.2, pp. 539 – 642.
Manum, S. B., Bose, M. N., Sawyer, R. T. 1991. Clitellate cocoons in
freshwater deposits since the Triassic. Zoologica Scripta
20:347 – 366.
Martin, A. J., Sealy, R. M. H., Young, J. O. 1994a. Does body size
differences in the leeches Glossiphonia complanata (L.) and
Helobdella stagnalis (L.) contribute to co-existence? Hydrobiologia 273:67 – 75.
Martin, A. J., Sealy, R. M. H., Young, J. O. 1994b. Food limitations
in lake-dwelling leeches: field experiments. Journal of Animal
Ecology 63:93 – 100.
Martin, A. J., Sealy, R. M. H., Young, J. O. 1994c. The consequences
of a food refuge collapse on a guild of lake-dwelling triclads
and leeches. Hydrobiologia 277:187 – 195.
Mather, C. K. 1954. Haemopis kingi, new species (Annelida:
Hirudinea). American Midland Naturalist 52:460 – 468.
Mather, C. K. 1963. Haemopis latero-maculatum, new species (Annelida: Hirudinea). American Midland Naturalist 70:168 – 174.
McLoughlin, N. J., Blinn, D. W., Davies, R. W. 1999. An energetic
evaluation of a predator – prey (leech-amphipod) couple in
Montezuma Well, Arizona, USA. Functional Ecology 13:45 – 50.
McNicol, D. K., Bendell, B. E., Mallory, M. L. 1995. Evaluating
macroinvertebrate responses to recovery from acidification in
small lakes in Ontario, Canada. Water, Air and Soil Pollution
85:451 – 456.
McNicol, D. K., Mallory, M. L., Mierle, G., Scheuhammer, A. M.,
Wong, A. K. H. 1997. Leeches as indicators of dietary mercury
exposure in non-piscivorous waterfowl in Central Ontario,
Canada. Environmental Pollution 95:177 – 181.
Meyer, M. C. 1940. A revision of the leeches (Piscicolidae) living on
freshwater fishes of North America. Transactions of the American Microscopical Society 59:354 – 376.
Meyer, M. C. 1946. A new leech Piscicola salmositica, n. sp. (Piscicolidae), from steelhead trout (Salmo gairdneri gairdneri
Richardson, 1838). Journal of Parasitology 32:467 – 476.
Meyer, M. C. 1975. A new leech, Macrobdella diplotertia sp. n.
(Hirudinea: Hirudinidae), from Missouri. Proceedings of the
Helminthological Society of Washington 42:83 – 85.
Meyer, M. C., Moore, J. P. 1954. Notes on Canadian leeches
(Hirudinea), with the description of a new species. Wasmann
Journal of Biology 12:63 – 96.
501
Milne, I. S., Calow, P. 1990. Costs and benefits of brooding in glossiphoniid leeches with special reference to hypoxia as a selection
pressure. Journal of Animal Ecology 59:41 – 56.
Misell, L. M., Shaw, B. K., Kristan W. B. Jr. 1998. Behavioral hierarchy in the medicinal leech, Hirudo medicinalis: feeding as a
dominant behavior. Behavioural Brain Research 90:13 – 21.
Moore, J. P. 1898. The leeches of the U.S. National Museum. Proceedings of the United States National Museum 21:543 – 563.
Moore, J. P. 1900. A description of Microbdella biannulata with
especial regard to the constitution of the leech somite. Proceedings of the Academy of Natural Sciences, Philadelphia
52:50 – 73.
Moore, J. P. 1901. The Hirudinea of Illinois. Illinois State Laboratory
of Natural History Bulletin 5:479 – 547.
Moore, J. P. 1906. Hirudinea and Oligochaeta collected in the Great
Lakes region. Bulletin of the Bureau of Fisheries 25:153 – 171.
Moore, J. P. 1912. Classification of the leeches of Minnesota, in: The
leeches of Minnesota. Geology and Natural History of Minnesota, Zoology Series No. 5:63150.
Moore, J. P. 1949. Hirudinea, in: R. Kenk, Ed. The animal life of
temporary and permanent ponds in southern Michigan. Miscellaneous Publications of the Museum of Zoology, University of
Michigan No. 71, pp. 38 – 39.
Moore, J. P. 1953. Three undescribed North American leeches
(Hirudinea: Notulae). National Academy of Natural Science,
Philadelphia No. 250, pp. 1 – 13.
Moore, J. P. 1954. Cited in Meyer and Moore (1954).
Moore, J. P. 1959. Hirudinea, in: Edmonson, W. T. Ed. Freshwater
Biology. 2nd ed. Wiley, New York, pp. 542 – 557.
Moore, J. P., Meyer, M. C. 1951. Leeches (Hirudinea) from Alaskan
and adjacent waters. Wasmann Journal of Biology 9:11 – 77.
Moser, W. E., Desser, S. S. 1996. Morphological, histochemical, and
ultrastructural characterization of the salivary glands and proboscises of three species of glossiphoniid leeches (Hirudinea:
Rhynchobdellida). Journal of Morphology 225:1 – 18.
Müller, F. 1844. De Hirudinea circa Berolinum hueusque observatis.
Dissitation Inaug., Berlin. pp 23 – 25.
Müller, O. F. 1776. Verrnium terrestrium et fluviatilum, seu animalium infusonorum helminthicorum, et testaceorum, non marinorum succincta historia. Hauniae et Lipsiae (Helminthica)
1:37 – 51.
Nehili, M., Ilk, C., Melhorn, H., Ryhnau, K., Dick, W., Njajou, M.
1994. Experiment on the possible role of leeches as vectors of
animal and human pathogens:a light and electron microscopy
study. Parasitology Research 80:277 – 290.
Nieczaj, R., Zerbst-Boroffka, I. 1993. Hyperosmotic acclamation in
the leech, Hirudo medicinalis L.: energy, metabolisms, osmotic,
conic and volume regulation. Comparative Biochemistry and
Physiology 106A:595 – 602.
Omodeo, P. 1998. History of Clitellata. Italian Journal of Zoology
65:51 – 73.
Oosthuizen, J. H., Davies, R. W. 1992. Redescription of the duck
leech, Theromyzon rude (Baird, 1869) (Rhynchobdellida: Glossiphoniidae). Canadian Journal of Zoology 70:2028 – 2033.
Oosthuizen, J. H., Davies, R. W. 1993. A new species of Theromyzon
(Rhynchobdellida: Glossiphoniidae), with a review of the genus
in North America. Canadian Journal of Zoology 71:1311 – 1318.
Pawlowski, L. K. 1948. Contribution a la connaissance des sangsus
(Hirudinea) de la Nouvelle-Ecosse, de Terre-Neuve et des iles
francais Saint-Pierre et Miquelon. Fragmenta faunistica Musei
Zoologici Polonici 5:317 – 353.
Pearse, A. S. 1932. Parasites of Japanese salamanders. Ecology
13:135 – 152.
Peterson, D. L. 1982. Management of ponds for bait leeches in
502
Ronald W. Davies and Fredric R. Govedich
Minnesota. Minnesota Department of Natural Resources, Section of Fisheries Investigation. Report No. 357:43.
Peterson, D. L. 1983. Life cycle and reproduction of Nephelopsis obscura Verrill (Hirudinea; Erpobdellidae) in permanent ponds of
northwestern Minnesota. Freshwater Invertebrate Biology
2:165 – 172.
Platt, T. R., Sever, D. M. Gonzalez, V. L. 1993. First report of the
predacious leech Helobdella stagnalis (Rhynchobdellida:Glossiphoniidae) as a parasite of an Amphibian, Ambystoma tigrinum
(Amphibia: Caudata). The American Midland Naturalist
129:208 – 210.
Prahacs, S. M., Hall, K. J. 1996. Leeches as in situ biomonitors of
chlorinated phenolic compounds. Part 1: Laboratory investigations. Water Research 30:2293 – 2300.
Prahacs, S. M., Hall, K. H. Duncan, W. 1996. Leeches as in situ biomonitors of chlorinated phenolic compounds. Part 2: Pulp mill
investigations. Water Research 30:2301 – 2308.
Pritchard, G. 1964. The prey of dragonfly larvae (Odonata:
Anisoptera) in ponds in northern Alberta. Canadian Journal of
Zoology 42:785 – 800.
Purschke, G., Westheide, W., Rohde, D. Brinkhurst, R. O. 1993.
Morphological reinvestigation and phylogenetic relationships of
Acanthobdella peledina (Annelida: Clitellata). Zoomorphology
(Berline) 113:91 – 101.
Pütter, A. 1907. Der Stoffwechsel der Blutegels (Hirudo medicinalis
L.). I. Zeitschriftfur Allgemeine Physiologie 6:217 – 286.
Pütter, A. 1908. Der Stoffwechsel der Blutegels (Hirudo medicinalis)
II. Zeitschrift für Allgemeine Physiologie 7: 16 – 61.
Qian, Y., Davies, R. W. 1994. Differences in bioenergetic and life-history traits between two generations of Nephelopsis obscura due
to the prehistory of their parents. Freshwater Ecology
8:102 – 109.
Qian, Y., Davies, R. W. 1996. Inter-population genetic variation in
the freshwater leech Nephelopsis obscura in relation to environmental heterogeneity. Hydrobiologia 325:131 – 136.
Rathke, H. 1862. Beitrage zur entwicklungsgeschicte der Hirudineen.
Leipzig, Verlag von Wihelm Engelmann 116 pp.
Reddy, D. C., Dratnal, E., Davies, R. W. 1992. Dynamics of energy
storage in a freshwater predator (Nephelopsis obscura) following winter stresses. Canadian Journal of Fisheries and Aquatic
Sciences 49:1583 – 1587.
Reddy, D. C., Davies, R. W. 1993a. Metabolic adaptations by the
leech Nephelopsis obscura during long-term anoxia and recovery. Journal of Experimental Zoology 265:224 – 230.
Reddy, D. C., Davies, R. W. 1993b. Bioenergetic responses to the initiation of winter and glycerol formation in the leech Nephelopsis obscura. Canadian Journal of Zoology 71:2036 – 2041.
Reynoldson, T. B., Bellamy, L. S. 1970. The establishment of interspecific competition in action between Polycelis nigra (Müll.)
and P. tenuis (Ijima) (Turbellaria, Tricladida). in: Proceedings of
the Advanced Study Institute, Dynamics Number Population,
Oosterbruck, pp. 282 – 297.
Reynoldson, T. B., Davies, R. W. 1976. A comparative study of the
osmoregulatory ability of three species of leech (Hirudinoidea)
and its relationship to their distribution in Alberta. Canadian
Journal of Zoology 54:1908 – 1911.
Reynoldson, T. B., Davies, R. W. 1980. A comparative study of
weight regulation in Nephelopsis obscura and Erpobdella punctata (Hirudinoidea). Comparative Biochemistry and Physiology
66:711 – 714.
Richardson, L. R. 1942. Observations on migratory behaviour of
leeches. Canadian Field Naturalist 56:67 – 70.
Richardson, L. R. 1969. A contribution to the systematics of the
hirudinid leeches with descriptions of new families genera and
species. Acta Zoologica Academiae Scientiarum Hungaricae
15:97 – 149.
Richardson, L. R. 1971. A new species from Mexico of the Nearctic
genus Percymoorensis and remarks on the family Haemopidae
(Hirudinoidea). Canadian Journal of Zoology 49:1095 – 1103.
Richardson, L. R. 1975. A contribution to the general zoology of the
land-leeches (Hirudinoidea: Haemadipsoidea Superfam. nov.).
Acta Zoologica Academiae Scientarium Hungaricae. 21:
119 – 152.
Riggs, M. R. 1980. Helminth parasites of leeches of the genus
Haemopis. Master’s thesis, Iowa State University.
Rosca, D. I. 1950. Duration of survival and variations in body
weight in Hirudo medicinalis placed in solutions of increasing
salinity. Studii si Cercetari de Biologie Cluj. 1:211 – 222.
Roters, F. J., Zebe, E. 1992a. Protease inhibitors in the alimentary tract
of the medicinal leech Hirudo medicinalis : in vivo and in vitro
studies. Journal of Comparative Physiology 162B:85 – 92.
Roters, F. J. Zebe, E. 1992b. Proteinors of the medicinal leech
Hirudo medicinalis:purification and partial characterisation of
three enzymes from the digestive tract. Comparative Biochemistry and Physiology 102B:627 – 634.
Sawyer, R. T. 1970. Observations on the natural history and behaviour of Erpobdella punctata (Leidy) (Annelida: Hirudinea).
American Midland Naturalist 83: 65 – 80.
Sawyer, R. T. 1972. North American freshwater leeches, exclusive of
the Piscicolidae, with a key to all species. Illinois Biological
Monograph No. 46,154 pp.
Sawyer, R. T. 1981. Leech biology and behavior. in: Muller, K. J.,
Nicholls, I. G., Stent, G. S. Eds. The neurobiology of the leech.
Cold Spring Harbor Lab., Cold Spring Harbor. New York,
pp. 7 – 26.
Sawyer, R. T. 1984. Arthropodization in the Hirudinea: evidence for
a phylogenetic link with insects and other Uniramia. Zoological
Journal of the Linnean Society, London 80:303 – 322.
Sawyer, R. T. 1986. Leech biology and behaviour. Clarendon, Oxford. 1065 pp.
Sawyer, R. T., Hammond, D. L. 1973. Distribution, ecology and behavior of the marine leech Calliobdella carolinensis (Annelida:
Hirudinea), parasitic on the Atlantic menhaden in epizootic
proportions. Biological Bulletin 145:373 – 388.
Sawyer, R. T., Shelley, R. H. 1976. New records and species of
leeches (Annelida: Hirudinea) from North and South Carolina.
Journal of Natural History 10: 65 – 97.
Sawyer, R. T., LePont, F., Stuart, D. K., Kramer, A. P. 1981. Growth
and reproduction of the giant glossiphoniid leech Haementaria
ghilianii. Biological Bulletin (Woods Hole, Mass.) 160:
322 – 331.
Say, T. 1824. Sur les Hirudo parasitica, laterialis, marmorata et decora dans le voyage du Major Long. Natural History, Zoology
Philadelphia 2:253 – 387.
Scudder, G. G. E., Mann, K. H. 1968. The leeches of some lakes in
the southern interior plateau region of British Columbia. Syesis
1:203 – 209.
Seymour, J. L., Henzel, W. J., Nevins, B., Stults, J. T., Lazarus, R. A.
1990. A potent glycoprotein IIb–IIIa antagonist and platelet
aggregation inhibitor from the leech Macrobdella decora. The
Journal of Biological Chemistry 265:10143 – 10147.
Shankland, M., Martindale, M. Q., Nardelli-Haefinger, D., Baxter,
E., Price D. J. 1992. Origin of segmental identity in the development of the leech nervious system. Development Supplement
2:29 – 38.
Siddall, M. E., Burreson, E. M. 1995. Phylogeny of the Euhirudinea:
independent evolution of blood feeding by leeches? Canadian
Journal of Zoology 73:1048 – 1064.
13. Annelida: Euhirudinea and Acanthobdellidae
Siddall, M. E., Burreson, E. M. 1996. Leeches (Oligochaeta Euhirudinea), their phylogeny and the evolution of life-history
strategies. Hydrobiologia 334:277 – 285.
Simon, T. W., Barnes, K. 1996. Olfaction and prey search in the carnivorous leech Haemopis marmorata. The Journal of Experimental Biology 199:2041 – 2051.
Singhal, R. N., Davies, R. W. 1985. Descriptions of the reproductive organs of Nephelopsis obscura and Erpobdella punctata
(Hirudinoidea: Erpobdellidae). Freshwater Invertebrate Biology
5:91 – 97.
Singhal, R. N., Davies, R. W. 1987. Histopathology of hyperoxia in
Nephelopsis obscura (Hirudinoidea: Erpobdellidae). Journal of
Invertebrate Pathology 50:33 – 39.
Singhal, R. N., Davies, R. W., Shah, K. L. 1985. The taxonomy and
morphology of the leeches (Hirudinoidea: Glossiphoniidae) parasitic on turtles from the Beas River (India) including descriptions of two new species and one redescription. Zoologischer
Anzeiger 215:147 – 155.
Singhal, R. N., Sarnat, H. B., Davies, R. W. 1988. Effect of anoxia
and hyperoxia on the neurons in the leech Nephelopsis obscura
(Erpobdellidae): ultrastructural studies. Journal of Invertebrate
Pathology 52:409 – 418.
Sladacek, V., Kosel, V. 1984. Indicator value of freshwater leeches
(Hirudinea) with a key to the determination of European
species. Acta Hydrochimie und Hydrobiologie 12:451 – 461.
Smiley, J. W., Sawyer, R. T. 1976. Osmoregulation in the leeches
Macrobdella dietra and Haemopsis marmorata. Bulletin of the
Association of Southeastern Biologists 23:96.
Smith, D. E. C., Davies, R. W. 1996a. Changes in energy allocation
by the predator Nephelopsis obscura exposed to differences in
prey availability. Canadian Journal of Zoology 75:606 – 612.
Smith, D. E. C. Davies, R. W. 1996b. Effects of aggregative behavior on the bioenergetics of the freshwater predatory leech
Nephelopsis obscura (Erpobdellidae). Freshwater Biology
36:647 – 659.
Soös, A. 1965. Identification key to the leech (Hirudinoidea) genera of
the world, with a catalogue of the species. I. Family: Piscicolidae.
Acta Zoologica Academiae Scientarium Hungaricae 2:417 – 463.
Soös, A. 1966a. Identification key to the leech (Hirudinoidea) genera
of the world, with a catalogue of the species. II. Families: Semiscolecidae, Trematobdellidae, Americobdellidae, Diestecostomatidae. Acta Zoologica Academiae Scientarium Hungaricae
12:145 – 160.
Soös, A. 1966b. Identification key to the leech (Hirudinoidea) genera
of the world, with a catalogue of the species. III. Family: Erpobdellidae. Acta Zoologica Academiae Scientarium Hungaricae
12:371 – 407.
Soös, A. 1966c. On the genus Glossiphonia Johnson, 1816, with a
key and catalogue to the species (Hirudinoidea: Glossiphoniidae). Annales Historico-Naturales Musei Nationalis Hungarici
58:271 – 279.
Soös, A. 1967. Identification key to the leech (Hirudinoidea) genera
of the world, with a catalogue of the species. IV. Family:
Haemodipsidae. Acta Zoologica Academiae Scientiarum Hungaricae 13:417 – 432.
Soös, A. 1968. Identification key to the species of the genus Erpobdella de Blainville,1818 (Hirudinoidea: Erpobdellidae). Annales
Historico-Naturales Musei Nationalis Hungarici 60:141 – 145.
Soös, A. 1969a. Identification key to the leech (Hirudinoidea) genera
of the world, with a catalogue of the species. V. Family: Hirudinidae. Acta Zoologica Academiae Scientiarum Hungaricae
15:151 – 201.
Soös, A. 1969b. Identification key to the leech (Hirudinoidea) genera
of the world, with a catalogue of the species. VI. Family: Glossi-
503
phoniidae. Acta Zoologica Academiae Scientiarum Hungaricae
15:397 – 454.
Spelling, S. M., Young, J. O. 1987. Predation on lake-dwelling
leeches (Annelida: Hirudinea): An evaluation by field experiment. Journal of Animal Ecology 56:131 – 146.
Treadwell, A. L. 1914. Polychaetous annelids of the Pacific coast in
the collections of the zoological museum of the University of
California. University of California Publications in Zoology
13:175 – 234.
Van Damme, N. 1974. Organogénèse de l’appareil genital chez la
sangsue Erpobdella octoculata L. (Hirudinée Pharyngobdelle).
Archives de Biologie 18:373 – 377.
Verrill, A. E. 1872. Descriptions of North American freshwater
leeches. American Journal of Science 3:126 – 139.
Verrill, A. E. 1874. Synopsis of the North American freshwater
leeches. Report to the United States Fisheries Commission
1872 / 73:666 – 689.
Von Brand, T. 1944. Occurrence of anaerobiosis among invertebrates. Biodynamica 4:185 – 328.
Webb, R.A. 1980. Spermatogenesis in leeches I. Evidence for a gonadotropic peptide hormone produced by the supraoesophageal
ganglion of Erpobdella octoculata. General and Comparative
Endocrinology 42:401 – 412.
Webb, R. A., Omar, F. E. 1981. Spermatogenesis in leeches. II. The
effect of the supraoesophageal ganglion and ventral nerve cord
ganglia on spermatogenesis in the North American medicinal
leech Macrobdella decora. Genetics and Comparative Endocronology 44:54 – 63.
Webster, H. 1879. Annelida Chaetopoda of New Jersey. New York
State Museum of Natural History Report 32:101 – 125.
Weisblat, D. A., Harper, G., Stent, G. S., Sawyer, R. T. 1980. Embryonic cell lineages in the nervous system of the glossiphoniid
leech Helobdella triserialis. Developmental Biology 76:58 – 78.
Whitman, C. O. 1886. The leeches of Japan. Quarterly Journal of
Microscopical Science 26:317 – 416.
Whitman, C. O. 1892. The metamerism of Clepsine. in: Festschrift
70 sten Geburtstag R. Leuckarts. 285 – 295.
Wicklum, D., Davies, R. W. 1996. The effects of chronic cadmium
stress on energy acquisition and allocation in a freshwater benthic invertebrate predator. Aquatic Toxicology 35:237 – 252.
Wicklum, D., Smith, D. E. C. Davies, R. W. 1997. Mortality, preference, avoidance, and activity of a predatory leech exposed to
cadmium. Archives of Environmental Contamination Toxicology 32:178 – 183.
Wilkialis, J., Davies, R. W. 1980a. The reproductive biology of
Theromyzon tessulatum (Glossiphoniidae: Hirudinoidea), with
comments on Theromyzon rude. Journal of Zoology (London)
192:421 – 429.
Wilkialis, J., Davies, R. W. 1980b. The population ecology of the
leech (Hirudinoidea: Glossiphoniidae) Theromyzon tessulatum.
Canadian Journal of Zoology 58:906 – 912.
Williamson, M. 1972. The analysis of biological populations.
Arnold, London.
Wrona, F. J., Davies, R. W. 1984. An improved flow-through
respirometer for aquatic macroinvertebrate bioenergetic research. Canadian Journal of Fisheries and Aquatic Sciences
41:380 – 385.
Wrona, F. J., Davies, R. W., Linton, L. 1979. Analysis of the food
niche of Glossiphonia complanata (Hirudinoidea: Glossiphoniidae). Canadian Journal of Zoology 57:2136 – 2142.
Wrona, F. J., Davies, R. W., Linton, L. Wilkialis, J. 1981. Competition and co-existence between Glossiphonia complanata and
Helobdella stagnalis (Glossiphoniidae: Hirudinoidea). Oecologia 48:133 – 137.
504
Ronald W. Davies and Fredric R. Govedich
Wrona, F. I., Linton, L. R., Davies, R. W. 1987. Reproductive success
and growth of two species of Erpobdellidae: the effects of water
temperature. Canadian Journal of Zoology 65:1253 – 1256.
Young, J. O. 1987. Predation on leeches in a weedy pond. Freshwater Biology 17: 161 – 167.
Young, J. O., Spelling, S. M. 1986. The incidence of predation on
lake-dwelling leeches. Freshwater Biology 16:465 – 477.
Young, S. R., Dedwylder, R., Friesen, W. O. 1981. Response of the
medicinal leech to water waves. Journal of Comparative Physiology 144: 111 – 116.
Young, J. O., Seaby, R. M. H., Martin, A. J. 1995. Contrasting mortality in young freshwater leeches and triclads. Oecologia
101:317 – 323.
Zaranko, D. T., Griffiths, R. W., Kaushik, N. K. 1997. Biomagnification of polychlorinated biphenyls through a riverine food web.
Environmental Toxicology and Chemistry 16:1463 – 1471.
Zebe, E., Roters, F. J., Kaiping, B. 1986. Metabolic changes in the
medicinal leech Hirudo medicinalis, following feeding. Comparative Biochemistry and Physiology 84A:49 – 55.
Zrzavy, J, Mihulka, S., Kepka, P., Bezdek, A., Tietz, D. 1998.
Phylogeny of the Metazoa based on morphological and 18S
ribosomal DNA evidence. Cladistics 14:249 – 285.
Zerbst-Boroffka, I. 1975. Function and ultrastructure of the nephridium in Hirudo medicinalis L. III. Mechanisms of the formation
of primary and final urine. Journal of Comparative Physiology
B 100:307 – 316.
14
BRYOZOANS
Timothy S. Wood
Department of Biological Sciences
Wright State University
Dayton, Ohio 45435
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Organ System Function
C. Environmental Physiology
III. Ecology and Evolution
A. Diversity and Distribution
B. Reproduction and Life History
C. Ecological Interactions
D. Evolutionary Relationships
IV. Entoprocta
V. Study Methods
VI. Taxonomic Key to North American
Freshwater Bryozoans
Literature Cited
I. INTRODUCTION
II. ANATOMY AND PHYSIOLOGY
Bryozoans are among the most commonly encountered animals that attach to submerged surfaces in
fresh water. During warm months of the year they are
found in almost any lake or stream where there are
suitable attachment sites. The variety of forms ranges
from wisps of stringy material to massive growths
weighing several kilograms. In the days before sand filtration, bryozoans were notorious for clogging the distribution pipelines of public water systems. Today they
still foul water intake lines, irrigation nozzles, and
wastewater treatment systems.
In general, bryozoans are sessile, modular invertebrates with ciliated tentacles that capture suspended
food particles. Historically they have included what are
now recognized as two very distinct and unrelated
phyla: Ectoprocta and Entoprocta. Within the ectoproct
bryozoans are two major classes: Phylactolaemata is an
exclusively freshwater group and the main focus of this
chapter; Gymnolaemata is a vastly larger, polyphyletic
collection of species, mostly marine except for a few
species in the subclass Ctenostomata which occur in
fresh or brackish water. In this chapter ctenostomes are
included in the general discussion. Entoproct bryozoans, phylogenetically distant from ectoprocts, are
treated in a separate section, but still included in this
chapter more for reasons of tradition than systematics.
A. External Morphology
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
1. Zooids and Lophophores
The phylactolaemate bryozoan colony is composed
of identical zooids fused seamlessly into a single structure (Fig. 1). Major anatomical features are shown in
Figure 2. Each zooid in the colony has two basic parts:
an organ system, or polypide, which can be partially
protruded as a unit; and a body wall which can enclose
the entire polypide and which separates the colony’s interior from the surrounding water. These parts are
joined in different places by three structures: a tentacular sheath, prominent retractor muscles, and a slender
funiculus loosely running from the gut caecum to a
point on the interior of the body wall.
The polypide bears a prominent lophophore composed of ciliated tentacles arranged around a central
mouth. In Fredericella species, the tentacles simply encircle the mouth (Fig. 3). In all other phylactolaemates
the lophophore extends dorsally in a bilateral pair
of arms, forming a U-shaped structure with an outer
row of long tentacles and an inner row of shorter ones
(Fig. 4). In most species, each tentacle bears one medial
and two lateral tracts of cilia which beat in metachronal
waves. Tentacles are loosely joined near their base by an
intertentacular membrane, effectively forming a groove
505
506
Timothy S. Wood
FIGURE 1 Two zooids of a tubular bryozoan colony, one retracted
and the other extended in a feeding position. Scale bar 0.5 mm.
between rows of tentacles. Two such grooves converge
from the lophophore arms toward the mouth. Adjacent
to mouth is the epistome, a small, heavily ciliated lobe
which may function in food selection.
In gymnolaemate (ctenostome) bryozoans the
zooids are tubular and rather delicate looking, with
small, conical lophophores (Fig. 8a – d). The body wall
is thin and transparent. Colonies are diffuse, consisting
of creeping chains of zooids sometimes separated by
narrow pseudostolons.
2. Statoblasts
A conspicuous feature of phylactolaemate bryozoans is the asexual production of encapsulated dormant buds, called statoblasts. The various distinctive
types are widely used for species identification. Most
species produce free statoblast that can be released
from the colony as disseminule, each bearing a periph-
eral band of sclerotized chambers normally filled with a
gas for buoyancy. In most species, the free statoblasts
(often called floatoblasts) acquire their buoyant gas in a
late developmental stage within the colony, while in a
few other species the floatoblast must be dried or
frozen before becoming buoyant. Many species with
self-inflating floatoblasts also produce specialized sessile statoblasts (“sessoblasts”) cemented to the substrate upon which colonies grow, and bearing a distinctive lamella around the rim. A third type of statoblast,
the piptoblast, is a simple, beanlike structure known
only in Fredericella species, lacking specialized external
structures and held firmly within the tubular branches
of the colony. Mukai (1982) provides an excellent
overview of statoblast development.
All statoblasts are composed of paired valves joined
at an equatorial suture. In most floatoblasts the valves
have two structural layers (Fig. 5a). An inner capsule
contains germinal tissue and food reserves; the outer
periblast covers the capsule completely and includes the
peripheral annulus and a central fenestra. In most
species, the so-called dorsal (or cystigenic) valve generally
has a smaller fenestra and correspondingly wider annulus
than the ventral (deutoplasmic) valve. At least one
species forms leptoblasts in which the capsule is lacking
and a fully formed zooid is enclosed by the periblast only
Sessoblasts have a similar arrangement of inner
capsule and outer periblast. The thin peripheral lamella
is considered homologous with the floatoblast annulus,
although it is never inflated. The basal valve provides a
large, irregular ring which adheres tightly to the substrate.
Gymnolaemate bryozoans do not produce statoblasts. Instead, many of the species restricted to fresh or
brackish water form special thick-walled hibernacula.
These irregularly shaped bodies are integrated into the
colony structure and attached firmly to the substrate,
remaining long after the colony disintegrates. Like statoblasts, hibernacula survive desiccation, changes of
salinity, and other suboptimal conditions, although the
limits of their resistance have not been documented.
B. Organ-System Function
1. Coelom, Neural System, and Body Wall
Most ectoproct colonies have a spacious coelom
shared by all zooids. A clear coelomic fluid is circulated
by cilia on the peritoneum. The main gastric coelom
communicates through an incomplete diaphragm with
the coelom of the lophophore, which extends fully into
every tentacle.
A small nerve ganglion is located on the diaphragm
at the base of the epistome between the mouth and the
14. Bryozoans
FIGURE 2
507
Schematic longitudinal section of a generalized colony (based on Mukai, 1982).
anus. A single nerve tract extends to each of the paired
arms of the lophophore, with nerves branching off to
each tentacle. Other nerves from the ganglion innervate
the epistome, the tentacle sheath, and the digestive tract.
There is no nervous communication between zooids.
The body wall is composed of living tissues, collectively called the endocyst, together with a nonliving
outer ectocyst (Fig. 2). Lining the main coelom is a peritoneum bearing scattered tracts of cilia. Behind the peritoneum lies a thin, longitudinal muscle layer followed
by a basement membrane. Overlying this is a thin layer
of circular muscles and a single layer of epidermis. Near
the apex of the zooid, the endocyst folds inward to
form a flat pocket, the vestibule, then joins the
FIGURE 3 Portion of a Fredericella colony showing characteristic circular outline
of the lophophore. Scale bar 2 mm.
508
Timothy S. Wood
FIGURE 4
Portion of a Lophopodella colony showing the typical horseshoe-shaped lophophore. Scale
bar 2 mm.
lophophore as an eversible tentacle sheath. When the
polypide is fully retracted, a well-defined opening, or
orifice, remains at the zooid tip. A pore in the vestibular
body wall of many species is normally closed except to
permit the release of floatoblasts from living colonies.
In some tubular species, a portion of the body wall
grows inwardly to form a kind of sclerotized, reinforc-
FIGURE 5
Sclerotized components of generalized phylactolaemate
statoblasts in exploded view. (a) Floatoblast; (b) sessoblast.
ing ring or incomplete septum. Those branches of tubular colonies that are attached to a substrate may have a
raised line, or keel, running along the outer surface.
The composition of the ectocyst is highly variable
among species. In most tubular colonies it is sclerotized
and remains intact for a time after living parts have
died. In at least two species, the ectocyst is composed
largely of chitinous microfibrils with random orientation. A thin, proximal electron-dense layer is overlain
by a thicker stratum of looser fibrils (Goethals et al.,
1984). Many organic and inorganic particles adhere to
this outer coat, including bacteria which also penetrate
the interior of the loose layer. The ectocyst in these
species varies from thin and flexible to leathery, opaque,
and brittle. With age the ectocyst may darken as the the
chitin is thickened and tanned. Thus, the growing tips
of the zooids are often distinctly lighter in color than
the rest of the colony. Rao et al. (1978) found that with
increasing age a second, separate sclerotized layer can
form in the inner body wall of Plumatella casmiana.
Mukai et al. (1984) have shown that the body wall
breaks down when contact is made between adhesive
pads from germinating statoblasts of the same species.
The resulting colonies are confluent, possibly allowing
a sort of outcrossing when sperm are circulated within
the common coelom.
In nontubular phylactolaemates the ectocyst is
gelatinous and often restricted to certain parts of the
14. Bryozoans
colony. Mukai and Oda (1980) reported many vacuolar cells, apparently with a secretory function, in the
epidermis of these species. The massive jellylike substance produced by Pectinatella magnifica is more than
99% water and contains partially denatured protein
along with some chitin, calcium, and sodium chloride
(Morse, 1930).
2. Feeding Mechanics
The lophophore is a complex feeding organ capable of capturing a wide variety of food particles. In
contrast with marine bryozoans, the lophophore of
most freshwater species is large and powerful, bearing
many closely positioned tentacles. Such an organ may
be particularly well suited to lentic habitats; certainly
those species with the largest lophophores seem to
grow best in quiet water. In species where lophophores
are tightly crowded (e.g., Cristatella or Pectinatella) the
processed water exits along defined excurrent channels.
This results in a significant feeding efficiency, according
to the model proposed by Eckman and Okamura
(1998). In a study using Plumatella repens, the ingestion rate per zooid was higher for large colonies than
for small ones, and also varied directly with ambient
water current (Okamura and Doolan, 1993).
Gilmour (1978) showed that differential beating of
the lateral and frontal cilia sort particles by density, rejecting heavier, inedible materials before they reach the
epistome. Kaminski (1991a) made a distinction between ciliary feeding, which brings in nannoplankton
(5 – 10 m), and tentacular feeding in which the tentacles orient segments of filamentous algae for ingestion.
Flicking movements of individual tentacles sometimes
throw particles toward the mouth; at other times the
tips of tentacles are brought together to prevent the escape of an active protozoan or rotifer.
Exactly how the lophophore captures minute particles is not entirely clear. In U-shaped lophophores the
membrane joining the base of all tentacles creates an
intertentacular groove of relatively still water. Bullivant
(1968) suggested that particles accelerating toward the
lophophore are impelled the final short distance into
this groove by their own momentum, while the water is
deflected abruptly to the sides. This would be consistent with the observation that species with the most
powerful lophophores are able to capture the smallest
particles (Kaminski, 1984).
Food particles entering the mouth collect momentarily in a short, ciliated pharynx, then are swallowed
through a narrow esophagus to the Y-shaped stomach.
A short unciliated cardia leads to the long, cylindrical
cecum, where particles are churned by rhythmic peristaltic contractions. In well-fed colonies waste products
collect in brownish, longitudinal bands, giving the ce-
509
cum a striped appearance. A short proximal part of the
cecum, the pylorus, leads through a narrow sphincter
to a straight intestine. Here processed particles are consolidated, infused with mucus, and ejected as a fecal
pellet through the anus, which lies just outside the
whorl of tentacles. The bundle of eliminated material is
too large to be re-ingested by neighboring zooids, and
it falls away from the colony.
3. Reproductive Systems and Larvae
Phylactolaemate bryozoans are sexually active during a single brief period of the year. Sperm develop in
conspicuous masses on the funiculus of certain zooids,
from which they later detach to circulate passively
throughout the colony coelom. Egg clusters develop on
the peritoneum ventrally within the zooid. There is
now direct evidence for outcrossing in populations of
Cristatella mucedo based on RADP assays (Jones et al.,
1994), but the mechanism for gamete transfer is unknown. Sperm motility has never been reported, nor
have sperm even been observed leaving one colony or
entering another. Self-fertilization thus appears likely in
most species except in instances where two colonies
have become confluent. Assuming that fertilization occurs in the coelom, the zygote somehow migrates to a
special sac formed as an ingrowth of the body wall.
The embryo develops into a specialized free-swimming
structure usually called a “larva,” although technically
a motile colony. Embryology of the planktonic larva
and the steps leading to its metamorphosis are summarized by Hyman (1959). Fine structure is described by
Franzén and Sensenbaugh (1983). The larva is composed of a heavily ciliated outer mantle and an inner
pear-shaped mass (Fig. 6a). The inner parts include one
to four fully formed polypides together with their funiculi and associated musculature. Released from the
colony usually after dark, the larva swims with its aboral pole forward, which contains special gland cells and
a neural center. Settling normally occurs within an hour
(Fig. 6b – d).
Among most Gymnolaemata, sexual reproduction
leads to the development of small larvae totally unlike
those of Phylactolaemata. Settling and metamorphosis
occur shortly after their release, since the larvae have
no feeding or digestive organs. The larva of Paludicella
and its embryological development were described
briefly by Braem (1896). However, virtually nothing is
known of larval ecology or behavior among freshwater
ctenostome bryozoans.
All bryozoan zooids are capable of budding new
zooids. In the Phylactolaemata, buds arise from a specific site on the midventral wall of the parental zooid
(Figs. 2 and 6a). The development of a new zooid is accompanied by the appearance of new bud primordia,
510
Timothy S. Wood
FIGURE 6 Settling and transformation of a larva. Scale bar for all figures 500 m. (a) Swimming larva
containing two fully developed zooids; the more tapered end is the oral pole; (b) the mantle peels back immediately after the larva attaches to a glass substrate; (c) constricted mantle appears as a dark mass; the two zooids
are now clearly visible, each with its own duplicate bud; (d) the mantle is drawn into the body cavity,
lophophores are extended, and the two zooids begin feeding (from Zimmerman, 1979).
which may or may not develop further. Brien (1953)
showed that every zooid bears two bud primordia: a
“main bud,” which forms the first daughter zooid, and
an “adventitious bud” between the main bud and
parental zooid, which becomes the second daughter
zooid. In addition, each main bud itself has a small
“duplicate bud” primordium on its ventral side. In the
process of budding, duplicate and adventitious buds
become main buds, and new duplicate and adventitious
buds are formed. These relations have been nicely clarified by Mukai et al. (1987).
In ctenostomes the new zooidal bud originates
from an outswelling of the body wall of a parental
zooid. An interior wall then grows across the base of
the bud to separate it from the parental body cavity.
Most often, a new bud develops at the distal end of a
lineal series of zooids, and as it grows a wall forms at
the tip to separate it from the body cavity of the next
distal bud in the series.
In addition to asexual budding, certain phylactolaemate species increase their numbers significantly by
an active fission of the colony. This is a common occurrence among the globular colonies of Cristatella, Pectinatella, Lophopodella, Lophopus, and Asajirella. The
products of fission glide slowly apart along the substrate by a means that is not fully understood.
C. Environmental Physiology
Little is known about the digestive physiology of
phylactolaemate bryozoans, except that the esophagus
environment is alkaline, the stomach is acidic, and the
intestinal pH is neutral (Marcus, 1926). Passage time
through the gut varies from one to 24 hs depending on
the ingestion rate. Zones of basophilic and acidophilic
cells line the cecum, but there is no agreement among
investigators on their function.
Statoblasts have been the subject of considerable
study. The period of obligate dormancy imposed on
most statoblasts serves to maintain populations
through periods of unfavorable environmental conditions. Those statoblasts not fixed to immobile substrates may also function in passive dispersal, carried
either by water currents or by migrating animals.
Brown (1933) demonstrated viability of Plumatella statoblasts after they had passed in separate trials through
the digestive tracts of a salamander, frog, turtle, and
duck.
Like statoblasts, the hibernacula of gymnolaemates
maintain the populations during periods of unfavorable conditions. Unlike most statoblasts, however, they
are an integral part of the colony, can never be released
freely into the water and thus would not normally
function as independent disseminules.
The resistance of statoblasts to environmental stress
has been reviewed by Bushnell and Rao (1974). Most
Plumatella and Fredericella statoblasts survive desiccation or freezing for periods of 1 to 2 years. Some Plumatella floatoblasts can germinate after more than 4
years in cold water. Lophopodella carteri statoblasts
have remained viable during dry storage for over 6
years. Other less hardy statoblasts include those of Pectinatella magnifica, which do not withstand desiccation
and survive only brief periods of freezing temperatures.
Statoblasts stored in water at a favorable temperature eventually germinate after a variable dormant
period. However, when that period is prolonged by
continuous exposure to cold, dry, anaerobic, or other
unfavorable conditions, a near simultaneous germination can be achieved when a suitable environment is
14. Bryozoans
restored. In some species, light is an important factor
for germination (Oda, 1980; Richelle et al., 1994). Ultrastructure of the statoblast suture and its possible relation to germination has been studied by Bushnell and
Rao (1974) and Rao and Bushnell (1979).
The budgeting of energy resources for colony
growth and reproduction is poorly understood. In
many cases, gametogenesis precedes statoblast production, but the two processes may also operate concurrently. The first zooid to germinate from a statoblast
never forms statoblasts itself, but it may participate in
gametogenesis. In laboratory colonies of Plumatella
emarginata, Mukai and Kobayashi (1988) found sexual activity only in certain colonies, which varied in
size from as few as 10 zooids. Only those colonies with
the most vigorous development of testes eventually
formed mature larvae.
The regulation of sessoblast development in Plumatellidae has not yet been clarified. Apparently, statoblast primordia can differentiate into either floatoblasts
or sessoblasts, and this differentiation becomes evident
during the late epidermal-disk stage (Mukai and
Kobayashi, 1988). In natural populations, Wood
(1973) noted that most sessoblasts were formed either
early or late in the season, leading him to suppose that
this was a response to suboptimal conditions. However, laboratory-reared Plumatella colonies frequently
form both floatoblasts and sessoblasts under apparently favorable growing conditions.
III. ECOLOGY AND EVOLUTION
A. Diversity and Distribution
Freshwater bryozoans are generally restricted to
relatively warm water, flourishing at 15 – 28°C. However, Cristatella mucedo has been found at 6°C (Bushnell, 1966), and Lophopus crystallinus survives at 0°C
for brief periods (Marcus, 1934). Fredericella indica
lives through the winter in most of North America,
budding new zooids at 3°C and producing piptoblasts
above 8°C. All of these species also grow well at higher
temperatures. Only Stephanella hina has never been
found above 17°C and is probably restricted to cool
water (Smith, 1989a). Temperatures as high as 37°C
were recorded for living Plumatella nitens and P. fruticosa (Bushnell, 1966). Shrivastava and Rao (1985)
noted that Plumatella emarginata in India shows only
meager growth at 34°C.
Phylactolaemates tolerate a wide range of pH, but
they favor slightly alkaline water. This was the conclusion of Tenney and Woolcott (1966) in a North Carolina study where pH ranged from 6.3 to 8.0. A Fred-
511
ericella species (probably F. indica) has been collected
in acidic conditions as low as pH 4.9 (Everitt, 1975),
Plumatella fruticosa at pH 5.7 (Bushnell, 1966), and
both P. repens and Hyalinella punctata at pH 6.3 (Tenney and Woolcott, 1966).
Most species occur in both still and running water,
but Plumatella emarginata and Fredericella indica grow
especially well in lotic habitats. Among those species
associated with still waters are Plumatella nitens, P. rugosa, P. recluse, Lophopus crystallinus, Hyalinella
punctata, and P. orbisperma. Many common species
tolerate turbid water, but Pectinatella magnifica does
not, possibly because its large colonies are necessarily
more exposed to settling particles (Cooper and Burris,
1984).
The reputation of bryozoans as clean water species
is not entirely warranted. Henry et al. (1989) found
Fredericella sultana, Plumatella emarginata, and P. fungosa thriving in high concentrations of heavy metals
and PCBs, although none of these had accumulated in
the tissues. Bushnell (1974) also recognized these
species to be particularly tolerant of extreme contamination from sewage and industrial wastes. More recently, Plumatella vaihiriae has been noted as an important fouling species in secondary clarifiers and
tertiary sand filters of wastewater treatment plants
(Wood and Marsh, 1999). According to Sládeček
(1980), both Plumatella repens and P. fungosa survive
at only 30% saturation of dissolved oxygen. Many
species, such as Lophopodella carteri flourish both in
the laboratory and the field when supplied with large
quantities of suspended organic particles. In Europe,
the distribution of Plumatella fungosa is correlated
with nutrient-enriched water (Job, 1976). Pectinatella
magnifica in the United States, Japan, and Korea become luxuriant in areas that are visibly eutrophic.
One clearly limiting factor for all species is the
availability of suitable surface on which to grow. Almost any solid, biologically inactive material is acceptable, including rocks, glass, plastics, automobile tires,
and aged wood. Surfaces that are seldom successfully
colonized include newly dead wood, corroded metal,
and oily or tarred materials. Aquatic plants are common substrates, although in a detailed survey of bryozoan – plant relations, Bushnell (1966) found little evidence for species specificity. Among large, floating
leafed plants, the lotus lily, Nelumbo lutea, supports
heavy growths of bryozoans, while other lilies, such as
Nymphaea or Nuphar, are seldom colonized.
Free floatoblasts, of course, are passive carriers of
living material to new substrates. Swimming larvae, on
the other hand, actively select attachment sites. Hubschman (1970) demonstrated particle size discrimination in larvae of Pectinatella magnifica, concluding that
512
Timothy S. Wood
rock particles smaller than 1-mm diameter were
avoided. Additional selection criteria are likely. Curry
et al. (1981) found evidence for selective settling of
Paludicella articulata larvae on mussel shells.
In general, freshwater bryozoan species are widely
distributed but variable in local abundance. Some
species appear to be distributed as metapopulations
with regular cycles of local extinction followed by new
immigration (Okamura, 1997). Two of our most common species, Fredericella indica and Plumatella reticulata, are rare in Europe (Massard et al., 1992; Massard
and Geimer, 1996); similarly, the abundant European
Plumatella fungosa is seldom found on our continent.
A near worldwide distribution is enjoyed by Plumatella
casmiana, Lophopodella carteri, and Paludicella articulata. However, the old notion that many other species
have a cosmopolitan distribution was apparently based
on faulty identification. Cristatella mucedo is restricted
to holarctic regions, while Fredericella browni and
Pottsiella erecta are most common where winters are
mild. About 25% of North American species are considered rare, and at least one, Lophopus crystallinus, is
seriously endangered. Some species are known to be
expanding their range: Pectinatella magnifica jumped
from North America to Europe and Japan and has recently invaded Korea; Lophopodella carteri is increasingly common since its introduction to North America
in the 1930s; and the once scarce Plumatella vaihiriae
is popping up in lakes and wastewater treatment facilities across the continent. Species believed to be endemic
to North America now include Plumatella nitens
(Wood, 1996), P. orbisperma (Ricciardi and Wood,
1992), P. recluse (Smith, 1992) and the tiny ctenostome, Sineportella forbesi (Wood and Marsh, 1996).
All known North American species have been reported east of the Mississippi River and north of the
39th parallel. Nineteen of the 24 species occur in states
bordering the Great Lakes and an additional two
species are described only from New England. Only the
brackish Victorella pavida has not been found further
north than Chesapeake Bay.
Relatively little is known about freshwater bryozoans west of Ontario and the Mississippi River. Most
states and provinces have no published records of bryozoans. However, various reports show Colorado and
Utah with natural populations of “Plumatella repens”,
P. casmiana, P. emarginata, P. fruticosa, P. fungosa,
Fredericella indica, and Cristatella mucedo. Other
western sightings show Cristatella mucedo in British
Columbia (Reynolds, 1976), and “Plumatella repens”
in Arizona and Nevada (Wilde et al., 1981; Dehdashti
and Blinn, 1986). Pectinatella magnifica appears to be
well established in eastern Texas (Neck and Fullington,
1983) and in the Pacific Northwest (personal commu-
nications). In Nebraska, the most abundant species appears to be Plumatella vaihiriae (Wood, unpublished).
The most recent regional surveys are those of Bushnell (1965a – c) in Michigan, Wood (1989) in Ohio,
Smith (1989b) in Massachusetts, Ricciardi and Reiswig
(1994) in eastern Canada, and Marsh and Wood in Illinois (in preparation). Each of these has revealed significant new species records for North America.
B. Reproduction and Life History
In most parts of the United States, bryozoan populations have two or more generations during the growing season. Only Fredericella indica typically is found
throughout the year living even under a cover of ice.
Populations of other species are normally maintained
during unfavorable seasons by dormant statoblasts. An
unusual exception to this pattern was reported by Dehdashti and Blinn (1986) in a population of “Plumatella repens” inhabiting an Arizona cave, where
nearly constant conditions permitted uninterrupted, active growth.
Winter dormancy in statoblasts is broken by conditions favorable to colony growth. In temperate climates
the principal triggering factor appears to be temperature. Viable statoblasts often germinate within a few
days of each other when the water temperature rises
above 8°C. In those regions where winter water temperature never approaches freezing, the time of germination may be more variable.
The single zooid that emerges from a statoblast is
termed the ancestrula. It eventually buds one-to-five
new zooids, each similarly capable of multiple zooid
budding. Colonies grow rapidly as water temperature
rises. Bushnell (1966) described naturally occurring
colonies of Plumatella nitens (identified at the time as
P. repens) which tripled and quadrupled in size within
one week. Doubling times of five days have been reported for both Plumatella casmiana and Fredericella
indica (Wood, 1973).
In species of Plumatella, Fredericella, and Lophopodella statoblasts may appear in colonies having
fewer than five zooids; in most other species they are
not formed until colonies are much larger. Among natural populations, a second generation usually arises either from the statoblasts or larvae of early spring
colonies, or even from fragments of colonies surviving
from the first period of growth. Third generation
colonies also are known, but any statoblasts they produce normally remain in diapause until the following
spring. Plumatella casmiana is unique in releasing thinwalled statoblasts which germinate immediately, so
there may be many overlapping generations throughout
the growing season.
14. Bryozoans
Gametogenesis, when it occurs, is normally encountered in the first spring generation of colonies, with larvae released 2 – 4 weeks after fertilization. Sperm masses
develop on the funiculus as ova appear on the peritoneum. The sperm later break free to circulate in the
coelom for 1 to 2 weeks. This is followed by larval development and eventual release. There appears to be little overlap in larval release dates among closely related
species living in the same area. In two Ohio lakes, freeswimming larvae of Plumatella rugosa appeared
throughout June; P. casmiana larvae were released during July; then came Hyalinella punctata in late
July – early August, followed by Plumatella emarginata
in late August – early September (Zimmerman, 1979).
Sexual activity in phylactolaemates is often reported as rare even in such careful studies as that of
Wöss (1996). Among the Plumatellidae, with a prolific
production of asexual statoblasts, sexual reproduction
would seem a relatively unimportant means for recruitment. However, in Pectinatella magnifica and Hyalinella punctata, with only one statoblast generation
per year, sexually produced larvae play a significant
role in establishing new colonies (Hubschman, 1970;
Wöss, 1999). Likewise, in Fredericella species, because
of their fixed statoblasts, larvae are assumed to be instrumental for both population growth and dispersal.
C. Ecological Interactions
The constant flow of water through bryozoan
lophophores creates a favorable environment for the
microscopic aufwuchs community. Protozoans, rotifers,
gastrotrichs, microcrustaceans, and other small animals
congregate especially on and around branching tubular
colonies of bryozoans. Flatworms, oligochaetes, snails,
orbatid mites, crayfish, and such insect larvae as caddisflies and midges occasionally graze on the living
zooids. Predation is seldom extensive.
Chironomid larvae enter old tubes of Plumatella
repens, hastening their disintegration, and occasionally
damaging living portions of the colony. Some workers
consider such damage to be accidental rather than the
result of direct feeding, while others regard chironomid
larvae as important predators. Colonies of P. casmiana
escape such harm, possibly because larvae cannot fit
themselves inside the narrow, branching tubules, but
instead build detritus tubes alongside the exterior
colony wall where they cause little harm. Pectinatella
magnifica is often a host to chironomid larvae, which
find shelter by burrowing into the gelatinous base close
to the substrate and causing no apparent damage. One
of these, Tendipes pectinatellae, is reported to be
specifically commensal on P. magnifica (Dendy and
Sublette, 1959).
513
Myxozoan and microsporidian parasites have long
been identified with freshwater bryozoan hosts. These
have been recently confirmed and further described
from Cristatella mucedo by Okamura (1996) and Canning et al. (1996, 1997). Myxozoans are easily seen
with light microscopy when they circulate in the
coelomic fluid. Infected colonies lose their normal vigor
and soon die. Buddenbrockia, an active, wormlike parasite, has been described from the coelom of several
plumatellids (e.g., Schröder, 1912; Marcus, 1941). Easily detected through the clear body wall of Hyalinella
punctata, for example, it seems to cause little distress
to the hosts as long as food remains abundant to the
host. Life cycle details of any of these parasites are so
far unknown.
Extensive predation by fish has not been verified.
Walburg et al. (1971) lists “bryozoans” among the stomach contents of six reservoir fish species, but provides no
details. When Dendy (1963) observed bluegills biting off
pieces of Plumatella sp., he believed the fish were doing
so only to obtain insect larvae associated with the
colonies. A homogenate of Lophopodella carteri tissues
is highly toxic to fish (Meacham and Woolcott, 1968),
and attempted feeding by fish on this or any other gelatinous species has never been observed. Freshwater prawns
will graze on colonies of Plumatella vaihiriae, but only
when no other food is available (Bailey-Brock and Hayward, 1984), suggesting either a lack of nutritional value
or a repellant chemistry of these colonies.
As sessile suspension feeders, bryozoans handle a
wide variety of food. Richelle et al. (1994) clearly
demonstrated the ability of Plumatella fungosa to
thrive on a diet of suspended bacteria. Other ingested
particles include diatoms, desmids, green algae,
cyanobacteria, dinoflagellates, rotifers, small nematodes, protozoa, and even microcrustaceans, along with
bits of detritus and inorganic materials. In a comparative study of three species, Kaminski (1984) found that
95% of ingested particles were under 5 m in diameter.
Plumatella repens, with its wide mouth and strong gut
musculature, ingested larger organisms than did either
Cristatella mucedo or Plumatella fruticosa. These included rotifers (Keratella sp.), colonial green algae, and
cyanobacteria as large as 75 m. In general, organisms
with long body extensions were able to avoid ingestion
by bryozoans, while small, rounded shapes were taken
most easily.
Analysis of stomach contents alone does not reveal
the important sources of bryozoan nutrition. Rotifers
and green algae have been known to pass through the
gut completely unharmed (Hyman, 1959) although
there are reports that a large portion of ingested organisms are variously damaged (Rüsche, 1938; Kaminski,
1991a).
514
Timothy S. Wood
The quantity of seston removed from a 460-ha eutrophic lake by Plumatella fungosa is estimated at 15
metric tons per year (Kaminski, 1991b). At the same
time, 8.8 tons of fecal pellets are deposited in the sediments, about twice the amount contributed by fish or
waterfowl, but less than that deposited by molluscs.
D. Evolutionary Relationships
Ectoprocta is one of three animal phyla collectively
known as lophophorates. The other two, Brachiopoda
and Phoronida, are represented by a small number of
solitary, mostly sessile, marine animals. For all of these,
the main unifying feature is a lophophore with hollow
tentacles containing an extension of the coelom, and
with cilia beating in metachronal waves to bring water
and suspended food toward the mouth.
As a group, the lophophorates show no clear affinity with any other invertebrates. The semblance of radial cleavage, mesoderm formation, and other elements
of morphology and development in branchiopods appears distinctly deuterostome (Zimmer, 1973; Emig,
1984). However, recent analysis of 18S rDNA sequence
data suggest that lophophorates are protostomes (Halanych et al., 1995), a conclusion supported by other
biochemical and morphological studies (e.g., Willmer,
1990; Gutmann, 1978).
Structural similarities have been noted between ectoprocts and the wormlike sipunculids. The anterior introvert of a sipunculid is protruded and withdrawn in a
manner very similar to that of the ectoproct polypide,
using coelomic pressure and retractor muscles. The
sipunculid introvert ends in a mouth surrounded by
hollow, ciliated, tentacular outgrowths. The gut is Ushaped, and the anus is situated near the base of the introvert.
In fact, nucleotide sequence data suggest that
sipunculids and gymnolaemate bryozoans may be
closely related, as are the brachiopods and phoronids
(Mackey et al., 1996; Cohen and Gawthrop, 1996).
However, the same data indicate that lophophorates, in
general, and ectoprocts, in particular, are not monophyletic assemblages. The gymnolaemate, Alcyonidium
gelatinosum and the phylactolaemate, Plumatella
repens appear to be neither closely related to each
other nor to the brachiopod – phoronid grouping.
Nevertheless, it is reasonable so far to propose an
ancestral line that led directly to phoronids and phylactolaemate ectoprocts. Only these two groups have a
crescentic lophophore and an epistome. They also both
produce new buds from a region on the oral (ventral)
side of the adult, while in other ectoprocts budding is
in an anal direction (Jebram, 1973). Many phoronids
produce special, fat bodies for energy storage, which
are strikingly similar to early developmental stages of
phylactolaemate statoblasts. While the larvae, in general, are quite different, the phoronid, Phoronis ovalis,
has a unique, ciliated actinotroch (Silén, 1954) that appears similar to the phylactolaemate “larva.” Significantly, P. ovalis also forms colonies which resemble
somewhat those of the phylactolaemate family Fredericellidae.
Phoronids and gymnolaemate ectoprocts are probably more distantly related, as reflected especially by
their different embryology and larval metamorphosis.
The phoronid actinotroch larva, for example, has a distinct coelom, the larval gut is retained in the adult, and
the lophophore develops from the metatroch ring of
cilia. By contrast, the gymnolaemate cyphonautes larvae has no larval coelom, the larval gut is not retained,
and tentacles develop from the episphere region.
Within the class Phylactolaemata, similarities in the
mode of colony growth and statoblast morphology
have long been regarded as the basis for evolutionary
relationships. Fredericella species are thought to exhibit
primitive features with their simple statoblasts and an
open, dendritic pattern of colony branching. The circular outline of the lophophore, associated with a relatively small number of tentacles, may also be a primitive character. The evolutionary trends include a shift
toward greater compactness of the colony and the accomodation of increasing numbers of tentacles on the
lophophore. Statoblasts serve the two seemingly conflicting roles of dispersal and of retaining a position on
proven favorable substrate.
Among the Plumatellidae and Stephanellidae all
species retain a basically tubular design. The two statoblast functions are served by distinctly different statoblast types: floatoblasts and sessoblasts.
In Lophopodidae, Pectinatellidae, and Cristatellidae the colonies are gelatinous and globular, with no
trace of branching. This development is coupled with a
larger lophophore and a single remarkable statoblast
design that incorporates a buoyant ring with marginal
hooks. These statoblasts function both as an anchor
and a means for dispersal. Radiating spines and hooks
were apparently acquired independently by all three
families in an interesting case of parallel evolution.
Hyalinella punctata, normally classified with the
Plumatellidae, shares many features with the Lophopodidae, and may actually link the two groups. Such features include a gelatinous colony wall, the absence of a
sessoblast, and a large floatoblast which, like those of
lophopodids, is initially nonbuoyant upon release.
Fossil statoblasts resembling those of recent Plumatella, Hyalinella, and Stephanella species appear in
rock strata from the Late Permian (Vinogradov, 1996),
demonstrating a remarkable stability in these taxa. Fu-
14. Bryozoans
ture fossil discovery and analysis could well provide
key information on phylactolaemate systematics.
The freshwater Gymnolaemata, all ctenostomes,
are members of an ancient marine group. They share
with the Cheilostome bryozoans many features expressed in the development of both larvae and colonies.
At this point, however, phylogenetic relationships are
still speculative.
IV. ENTOPROCTA
Entoprocta is a small group of about 60 species
distinct in almost every way from the Ectoprocta but
historically included with them under the name “bryozoan.” Both groups are sessile with ciliated tentacles
and an incomplete separation of budded zooids, but
the similarities stop there. The only known freshwater
species, Urnatella gracilis, was discovered in North
America in 1851. Two other species have been proposed elsewhere based mainly on number of stalk segments and morphology of the basal plate. Emscher-
FIGURE 7
515
mann (1965) considers all of these synonymous with
U. gracilis. Although normally classified as Urnatellidae, the genus is similar to the marine species Barentsia
and may be united with it in the family Pedicellinidae.
The zooid is a bulbous calyx borne on a flexible,
segmented stalk measuring up to 5-mm long (Fig. 7).
Several such stalks, either solitary or sparingly
branched, may arise from a basal plate. The calyx
bears a single whorl of 8 – 16 uniformly short, ciliated
tentacles. The area of the calyx enclosed by the tentacles, called an atrium or vestibule, includes both the
mouth and the anus. When the zooid is disturbed, the
tentacles fold over the vestibule and are covered by a
tentacular membrane.
Internal organs include a fully ciliated digestive
tract and a large medial ganglion. A number of excretory flame bulbs communicate through short ducts to a
common nephridiopore; additional flame bulbs occur
in the stalk. The body cavity is a pseudocoel filled with
loose mesenchyme and extending into each tentacle.
The calyx is deciduous, dropping off at the onset of
cold temperatures or other unfavorable conditions. The
Small entoproct colony (Urnatella gracilis) showing external structures.
516
Timothy S. Wood
basal segments of stalks, containing food and germinal
tissue, can survive the winter much like ectoproct statoblasts, forming new calyxes when the water temperature returns to around 15°C. New zooids develop by
budding from the stalk, as illustrated particularly well
by Oda (1982). It is reported that young colonies can
disperse locally by crawling over the substrate (Protosov, 1980). Sexual reproduction leads to the development of swimming larvae, presumably similar to those
of other Pedicellinidae; details are unknown.
Like other bryozoans, Urnatella gracilis is a suspension feeder, consuming organic particles, unicellular
algae, and protozoans. Two especially important foods
are the diatom Melosira and two green algae species,
Pediastrum duplex and P. simplex (Weise, 1961).
Urnatella gracilis is known from every continent
except Antarctica and Australia. In North America its
reported distribution ranges from the east to the west
coast and from Florida, Louisiana, and Texas to as far
north as Michigan. Zooids attach to almost any substrate, including rocks, sticks, aquatic plants, bivalve
shells, ectoprocts, and such debris as nails, beverage
cans, and lead fishing weights (Eng, 1977). Individuals
occur most frequently in flowing water or in shallow
areas of large lakes where there is extensive water
movement. Tolerance to a wide range of chemical and
physical conditions has been noted most recently by
Hull et al. (1980), and Cusak and McCullough (1985).
In some areas, especially on new substrate,
entoprocts may comprise a sizeable fraction of the
macroinvertebrate community. Stalk densities of over
225,000 / m2 and an average biomass of 868 mg / m2
(dry weight) were reported from a seven month old artificial stream in Mississippi (King et al., 1988).
Entoproct phylogeny is even less clear than that of
the Ectoprocta. Any similarities to ectoproct bryozoans
are clearly the result of convergent evolution rather than
common ancestry. A close comparison of entoproct tentacles and the ectoproct lophophore reveals little in common despite their outwardly similar structure and function. Body cavities of the two groups are likewise
irreconcilable: a pseudocoel on one hand and a true
coelom on the other. Analysis of 18S r-RNA data support
a wide separation between entoprocts and ectoprocts,
placing the entoprocts instead among the pseudocoelomates (Mackey et al., 1996). Nevertheless, puzzles such
as this are expected in animals so highly modified by sessile life style and modular architecture, and for now the
question of entoproct origins remains open.
V. STUDY METHODS
Bryozoans are normally found in areas of shallow
water where there is a suitable firm substrate. Although
most species are plainly visible to the unaided eye, a
10 Coddington-type magnifier is useful for field identification. Many species can be detected by examining
the edges of floating objects for their free statoblasts.
Some gelatinous colonies can be nudged gently from
the substrate with little damage, but with other species
it is better to collect pieces of substrate with colonies
attached. Basic equipment for this includes a sturdy
knife for organic substrates and a cold chisel and hammer for rocks. In lakes, where colonies are elusive, statoblasts can often be retrieved from sediments using a
stack of standard seives with mesh openings of 1.0
mm, 500, and 150 m (Jones et al., 1998).
Swimming larvae are most easily detected by placing large colonies in a shallow tray of water. If larvae
are present, they will generally be released with gentle
prodding, or will emerge spontaneously during the
nighttime hours. They are best seen against a dark
background. Before their release from the colony,
zooids with larvae appear swollen at the tip.
Bryozoans are not difficult to rear in the laboratory, but the substrate on which they attach must always be inverted so that fecal material and other settling debris will not accumulate around the zooids.
Most species can be kept at room temperature. At
lower temperatures there are fewer problems with fouling organisms, but colony growth is slow. Food appears to be the most important variable for success
with laboratory-reared bryozoans. Oda (1980) maintained colonies of Lophopodella carteri on pure cultures of Chlamydomonas reinhardtii. Wayss (1968)
kept Plumatella repens for three years using a variety
of unicellular green algae in 1:1 Knop’s solution and
soil extract. Others have achieved limited short-term
success using mixed protozoan cultures, pet fish food,
and fine detritus. One reliable and trouble-free source
of food is simply the suspended organic particles circulated through a dark rearing tank from a large, well-lit
aquarium in which active fish are maintained (Wood,
1996).
If specimens are to be preserved, they should be
anaesthetized before fixing so the lophophores will remain extended. The most convenient method is to confine colonies to a small covered dish of water with thin
wafers of menthol floating on the surface. The menthol
diffuses slowly into the water and relaxes the zooids
within an hour or two. Lophophores of anaesthetized
zooids can be hardened with drops of full-strength formalin. The colony is then fixed and preserved in 70%
ethyl alcohol.
Confirmed identification of most species requires
the presence of statoblasts inside the colony. Statoblast
dimensions and surface topography are species-specific.
In specimens stored for an appreciable time, any gas in
the floatoblast annulus is replaced by liquid, making
14. Bryozoans
the entire capsule clearly visible through the periblast.
In this case, it is important not to mistake capsule dimensions for those of the fenestra. Length and width of
sessoblasts should be measured from the base of the
lamella, not from its outer edge.
Surface features of the floatoblast fenestra can often be seen in isolated valves using ordinary light microscopy. However, features of the annulus require the
use of a scanning electron microscope. The most common surface pattern in the floatoblast annulus shows
the sclerotized cells in relief, like cobblestones, a condition called “paved” (Fig. 12a). The annulus may also
be reticulated (Fig. 12b) or adorned with tiny nodules
(Fig. 12d). In various species the floatoblast fenestra
may be smooth (Fig. 12a), reticulated (Fig. 12b), tuberculated, or bearing a reticulum in which each cell contains a single interstitial tubercle (Fig. 12d). Other diagnostic features of the floatoblast include polar grooves
on the dorsal valve (Fig. 11b) and appearance of the intact suture (Fig. 12c). All recent species descriptions include statoblast details such as these.
For detailed examination of a statoblast using light
microscopy, it is useful to first separate the component
parts by placing it for about 1 min in a hot solution of
potassium hydroxide. Then transfer it to distilled water
where the valves should separate spontaneously. Use
fine insect pins (size 000) to assist in this process and to
517
tease away the yolky contents. I find it convenient to
arrange these parts under the microscope and photograph them for my working records.
To prepare statoblasts for scanning electron microscopy it is first necessary to remove manually the
thin membrane that often adheres tightly to the surface. If subsequent cleaning is necessary, ultrasonic
treatment is too harsh. Instead, place the statoblasts in
a small Eppendorf vial with a 0.05 M sodium hexametaphosphate, then hold the vial against the shaft of an
electric vibrating blade shaver. Wash the statoblasts in
several changes of distilled water. Freeze-drying helps
prevents distortion of the fenestra. Sputtering is advisable but not essential.
Well over half the recognized phylactolaemate
species, are represented by type specimens in major
museums, including all species named since the 1940s
(Wood, 1999). Even long neglected and desiccated material remains valuable for morphological study of statoblasts.
Certain techniques for subcellular study of freshwater bryozoans have proven particularly useful. These
include analysis by karyotype (Backus and Mukai,
1987), by randomly amplified polymorphic DNA
(Okamura et al., 1993), 18S ribosomal DNA (Halanych et al., 1995), and the identification of polymorphic microsatillite loci (Freeland et al., 1999).
VI. TAXONOMIC KEY TO NORTH AMERICAN FRESHWATER BRYOZOANS
1.
Zooid composed of bulbous body on externally segmented stalk; tentacles folding individually toward center when zooid is disturbed; statoblasts absent (Fig. 7). Phylum Entoprocta . . . Urnatella gracilis Leidy, 1851.
1.
Zooid without externally segmented stalk; tentacles withdrawn together when zooid is disturbed; statoblasts may be present. Phylum Ectoprocta...................................................................................................................................................................................2
2(1).
Colony composed of branching tubules, body wall transparent-to-opaque; statoblasts (if present) with smooth margins; tentacles
fewer than 65 .....................................................................................................................................................................................3
2.
Colony globular and either lobed or entire in outline; body wall transparent; statoblasts with peripheral spines, hooks, or pointed
extensions; sessoblasts absent; tentacles more than 65 .....................................................................................................................25
3(2).
Extended lophophore circular in outline; statoblasts (if present) piptoblasts only (Fig. 9a, b); tentacles fewer than 25.......................4
3.
Extended lophophore U-shaped in outline; statoblasts either floatoblasts or sessoblasts (or both); tentacles more than 25 ..............10
4(3).
Statoblasts never formed; ectocyst stiff, shiny, and transparent; tentacles fewer than 20; individual zooids clearly demarcated by internal septa; orifice appears quadrangular when lophophore is withdrawn. Gymnolaemata, Ctenostomata.......................................5
4.
Statoblasts formed, especially in zooids attached to substrate, but possibly missing in some specimens; ectocyst not stiff and shiny;
tentacles more than 19. Phylactolaemata, Fredericellidae, Fredericella ...............................................................................................8
5(4).
Each zooid includes a uniformly narrow, sinuous, stolonlike tubule by which it is joined to a parental zooid ....................................6
5.
Stolonlike tubules absent, zooids branch from each other at nearly right angles; widely distributed in North America and worldwide. (Fig. 8b). Paludicellidae. Paludicella articulata (Ehrenberg, 1831).
6(5).
Tentacles numbering exactly 8. Victorellidae ......................................................................................................................................7
6
Tentacles more than 15; zooids ranging from straight and erect-to-bulbous and recumbant. (Fig. 8c). Known throughout eastern
North America, especially south of the 40th parallel. Pottsiellidae. Pottsiella erecta (Potts, 1884).
518
Timothy S. Wood
Gymnolaemate colonies occurring in fresh water. Scale bar for all figures 1 mm. (a) Sineportella
forbesi; (b) Paludicella articulata; (c) Pottsiella erecta; and (d) Victorella pavida.
FIGURE 8
7(6)
Erect portion of zooid 0.6 – 1.6 mm long and only slightly contracile (Fig. 8d); occurring mainly in brackish water; reported in
North America south from Chesapeake Bay and in southeastern Louisiana. Victorella pavida (Kent, 1870).
7
Erect portion of zooid never more than 0.3 mm long and highly contractile (Fig. 8a), known only from a single site in east central
Illinois. Sineportella forbesi Wood and Marsh, 1996.
8(4)
Piptoblast surface densely pitted or minutely roughened; appearing dull and granular when dry........................................................9
8
Piptoblast surface appearing mirror-smooth and shiny when dry. Common in Europe, confirmed in North America only from a single site in Curry County, Oregon. Fredericella sultana (Blumenbach, 1779).
9(8)
Piptoblast broadly oval to round (Fig. 9b), often more than one per zooid; valves covered by a minutely wrinkled mantle which is
easily removed by 5-min immersion in concentrated KOH solution to reveal smooth sclerotized surface. Reported throughout
North, Central and South America. Fredericella browni (Rogick, 1945).
9
Piptoblast oval-to-elongate (Fig. 9a), seldom more than one per zooid; surface uniformly pitted and unaffected by KOH. Common
throughout North America, also scattered sites in Europe and Asia; probably includes several species not yet distinguished. (Fig.
13a). Fredericella indica Annandale, 1909.
10(3)
Floatoblasts circular or nearly so, with little difference in appearance between dorsal and ventral sides; colony wall thick, soft,
transparent. Uncommon...................................................................................................................................................................11
10
Floatoblasts oval-or-oblong; fenestrae of two valves distinctly different in size; colony wall not necessarily thick or transparent.
Abundant and widespread. Plumatellidae 13
11(10)
Zooids erect, extending up to 4 mm. but collapsing when removed from water; polypide and lophophore relatively small; floatoblast fully reticulate and lacking tubercles (Fig 9m). Confirmed in North America only from several sites in Massachusetts, although
floatoblasts reported from Oregon. Stephanellidae. Stephanella hina Oka 1908.
14. Bryozoans
519
Free statoblasts from the families Plumatellidae and Fredericellidae. Scale bar for all figures 250
m. (a) Fredericella indica; (b) Fredericella browni; (c) Plumatella casmiana (capsuled floatoblast); (d) Plumatella casmiana (leptoblast); (e) Plumatella repens; (f) Plumatella fruticosa; (g) Plumatella reticulata; (h) Plumatella emarginata; (i) Hyalinella punctata; (j) Plumatella vaihiriae; (k) Plumatella orbisperma; (l) Plumatella
bushnelli; and (m) Stephanella hina [a – e and g – i modified from Wood, 1989; j based on Rogick and Brown,
1942; k and m based on Bushnell, 1965b].
FIGURE 9
11
Zooids not as above, floatoblast surface variable. Plumatellidae ......................................................................................................12
12(11)
Floatoblast fenestra tuberculated, annulus adorned with minute nodules (Figs. 12d and 9k), sessoblast circular or nearly so; thick
walls of large colonies fusing into a firm, transparent matrix. Known only from a few sites in Michigan and the northern Great
Lakes. Plumatella orbisperma (Kellicott, 1882).
12
Floatoblast fully reticulated and without tubercles; sessoblast long oval-to-rectangular. Known only from small ponds in forested
sites of New England. Plumatella recluse Smith, 1992.
13(10)
Colony wall thick, soft, and transparent; colony entirely adherant to substrate; zooids forming low mounds when polypides are retracted (Fig. 13d); floatoblast width greater than 300 m, length over 400 m (Fig. 9i), dark and seldom buoyant upon release
from colony, sessoblasts never formed. Widely distributed. Hyalinella punctata (Hancock, 1850).
13
Floatoblast and colony not as above, or sessoblasts present .............................................................................................................14
14(13)
Width of floatoblast dorsal annulus at poles never less than length of dorsal fenestra (Figs. 14.9g, h); internal septa frequent.........15
14.
Width of floatoblast dorsal annulus at poles less than length of dorsal fenestra (Fig. 9e, f, j, l); internal septa present or absent. .....16
15(14).
Floatoblast dorsal valve nearly flat, long edge curved; suture between valves visible in dorsal view (Figs. 5a, 9h, and 12a); sessoblast
surface uniformly granular, tentacles about 40. Internal septa perpendicular to colony wall; mouth region often red. Common and
widely distributed, especially in flowing water. Plumatella emarginata Allman, 1844.
15.
Floatoblast valves almost equally convex, long edge relatively straight (Fig. 9g), sessoblast surface roughened by network of raised
lines; tentacles about 32; internal septa slightly oblique; mouth region never red. Common throughout North America and south at
least to Panama, also reported from Israel. Plumatella reticulata Wood, 1988.
520
Timothy S. Wood
FIGURE 10 Statoblasts in the families Lophopodidae, Pectinatellidae, and Cristatellidae. Scale bar for all figures 500 m. (a) Pectinatella magnifica; (b) Cristatella magnifica; (c) Lophopodella carteri; (d) Lophopus
crystallinus [a – c modified from Wood, 1989].
16(14).
Floatolast and sessoblast length not more than twice width .............................................................................................................17
16.
Floatoblast and sessoblast length more than twice width (Fig. 9f); colonies with thin branches largely free of substrate. Widely distributed, especially in northern, oligotrophic habitats. Plumatella fruticosa Allman, 1844.
17(16)
Floatoblast length-to-width greater than 1.6 (Fig. 9c, d); colony composed of short, profusely branched tubes always closely adherent to substrate (Fig. 13b), tubes becoming erect and fused when crowded; floatoblast dorsal fenestra lacking prominent tubercles;
Scanning electron micrographs showing floatoblast features. Scale bar mm. (a) Lateral view;
(b) frontal view of dorsal valve. cp, central prominance; da, dorsal annulus; df, dorsal fenestra; pg, polar
groove; su, suture; and vf, ventral fenestra. Scale bar 100 m.
FIGURE 11
14. Bryozoans
521
delicate thin-walled floatoblast sometimes present (Fig. 9d); sessoblast lamella relatively narrow. Common and widely distributed.
Plumatella casmiana (Oka, 1907).
17
Floatoblast length to width less than 1.6 (Fig. 9e,j,l); colony branches not particularly short, colony or statoblasts not as above ....18
18(17)
Floatoblast distinctly tapered at the poles, with strongly reticulated fenestrae (Figs. 9j, 12b); dorsal annulus bulging around inner
capsule to form a distinct shoulder; sessoblast surface densely pitted and without tubercles. Uncommon but often locally abundant,
ranging throughout the United States. Plumatella vaihiriae (Hastings, 1929).
18
Floatoblast rounded at poles, fenestra not strongly reticulated, annulus extending evenly from suture to fenestra without bulging
around capsule; sessoblast surface densely tuberculate .....................................................................................................................19
19(18)
Floatoblast lateral edges relatively straight (Fig. 9l); dorsal fenestra with prominent tubercles annulus with dense nodules visible by
SEM; sessoblast lamella relatively wide. Known only from a single site on a North Carolina barrier island. Plumatella bushnelli
Wood, 2001.
19
Floatoblast lateral edges evenly curved (Fig. 9e); dorsal fenestra with weak tubercles and slight reticulation; common and widespread ..............................................................................................................................................................................................20
20(19)
Floatoblast dorsal fenestra small, not more than half the diameter of ventral fenestra; ventral annulus markedly wider at poles than
at sides .............................................................................................................................................................................................21
20
Floatoblast dorsal fenestra large, considerably more than half the diameter of ventral fenestra (Fig. 11b); ventral annulus uniformly
narrow. Known from Massachusetts to Wisconsin, mostly north of the 41st parallel. Plumatella nitens Wood, 1996.
FIGURE 12 Scanning electron micrographs showing floatoblast surface morphology. Scale bar mm. (a)
Plumatella emarginata dorsal valve with “paved” annulus and smooth fenestra; (b) Plumatella vaihiriae with
reticulated fenestra and annulus; (c) Hyalinella punctata with paved annulus and suture as a raised cord; and (d)
Plumatella orbisperma with reticulation and “interstitial” tuibercles on the annulus, small nodules on fenestra.
522
Timothy S. Wood
21(20)
Colony tubules seldom fused together; scanning electron microscopy (SEM) reveals no tubercles on floatoblast annulus, although
tiny nodules may be present, internal septa are uncommon. .............................................................................................................22
21
Large colony forming tight masses of fused tubules without free branches; SEM reveals tubercles on floatoblast annulus; many
internal septa; occurring in highly eutrophic water; abundant in Europe, but generally uncommon in North America. Plumatella
fungosa (Pallas, 1768).
22(21)
SEM shows floatoblast annulus surface smooth except for tiny, rash-like nodules............................................................................23
22
SEM shows floatoblast annulus surface roughened by concave cell walls, nodules absent; species common and widespread in North
America, especially in lentic habitats (Fig. 13c). Plumatella rugosa (Wood, Geimer and Massard 1998).
23(22)
SEM shows nodules only on floatoblast annulus ..............................................................................................................................24
23
SEM shows nodules abundant on both floatoblast annulus and fenestra; known from Illinois, Ohio, and western New York.
Plumatella nodulosa Wood, 2001.
24(23)
Floatoblast suture defined by a lumpy, double cord; known only from Illinois. Plumatella similirepens Wood, 2001.
24
Floatoblast suture marked by two crooked rows of large, toothlike tubercles; known from Europe and New Zealand, but not yet
documented from North America. Plumatella repens (Linnaeus, 1758)
25(2)
Mouth region with red pigmentation; prominent pair of white spots at end of each arm of lophophore; statoblasts round with
hooked spines radiating from outer margin of annulus (Fig. 10a). Colony gelatinous and slimy, ranging from a flat sheet to footballsized mass. (Fig. 13f). Common and widely distributed. Pectinatellidae. Pectinatella magnifica (Leidy, 1851)
FIGURE 13
Various colony forms in Phylactolaemata. (a) Fredericella indica growing on a piece of wood,
showing many free branches, scale bar 5 cm; (b) Plumatella casmiana, with zooids attached throughout
their length to the substrate or to each other, scale bar 2 mm; c) Plumatella rugosa growing on the inner
surface of a mussel valve, scale bar 2 cm; (d) Hyalinella punctata, showing both densely packed and freeranging growth, all zooids firmly attached to the substrate, scale bar 5 mm; (e) Cristatella mucedo, showing
a single statoblast inside, scale bar 5 mm; (f) Pectinatella magnifica growing on the underside of a small log,
scale bar 2 cm [b from Rogick, 1941; c from Rogick and Brown, 1942; c– f from Wood, 1989].
14. Bryozoans
523
25
Mouth region without red pigmentation, lophophore lacking pair of white spots ............................................................................26
26(25)
Colony not distinctly linear, statoblasts oblong, not round...............................................................................................................27
26
Colony linear, often longer than 2 cm (Fig. 13e); statoblasts round with wiry, hooked spines radiating beyond periphery from margin of fenestrae of both valves (Fig. 10b). Occuring mainly in oligotrophic waters. Cristatellidae. Cristatella mucedo Cuvier, 1798.
27(26)
Statoblast with series of small hooks localized along the polar margins (Fig. 10c). Uncommon, but can be locally abundant (Fig. 4).
Lophopodella carteri (Hyatt, 1866).
27
Statoblast tapering to a single point at each pole (Fig. 10d); extremely rare worldwide, and possibly endangered. Lophopus crystallinus (Pallas, 1768).
*
Species dropped from an earlier version of this key include Stolella indica, S. evelinae, and Plumatella coralloides. Recent work
throws doubt on their reported occurrence in North America (Wood, 1998).
LITERATURE CITED
Backus, B. T. Mukai, H. 1987. Chromosomal heteromorphism in a
Japanese population of Pectinatella gelatinosa and karyotypic
comparison with some other phylactolaemate bryozoans. Genetica 73:189 – 196.
Bailey-Brock, J. H., Hayward, P. J. 1984. A freshwater bryozoan,
Hyalinella vaihiriae Hastings (1929), from Hawaiian prawn
ponds. Pacific Science 38:199 – 204.
Braem, F. 1896. Die geschlectliche Entwicklung von Paludicella
Ehrenbergii. Zoologischer Anzeiger 19:54 – 57.
Brien, P. 1953. Etude sur les Phylactolémates. Annales de la Société
Royale Zoologique de Belgique 84:301 – 444.
Brown, C. J. D. 1933. A limnological study of certain fresh-water
Polyzoa with special reference to their statoblasts. Transactions
of the american Microscopical Society 52:271 – 313.
Bullivant, J. S. 1968. The rate of feeding of the bryozoan,
Zoobotryon verticillatum. New Zealand Journal of Marine and
Freshwater Research 2:111 – 134.
Bushnell, J. H. 1965a. On the taxonomy and distribution of freshwater Ectoprocta in Michigan. Part I. Transactions of the
American Microscopical Society 84:231 – 244.
Bushnell, J. H. 1965b. On the taxonomy and distribution of freshwater Ectoprocta in Michigan. Part II. Transactions of the
American Microscopical Society 84:339 – 358.
Bushnell, J. H. 1965c. On the taxonomy and distribution of freshwater Ectoprocta in Michigan. Part III. Transactions of the American Microscopical Society 84:529 – 548.
Bushnell, J. H. 1966. Environmental relations of Michigan Ectoprocta, and dynamics of natural populations of Plumatella
repens. Ecological Monographs 36:95 – 123.
Bushnell, J. H. 1974. Bryozoa (Ectoprocta), in: Hart, C. W. Fuller, S.
L. H. Eds., Pollution ecology of freshwater invertebrates. Academic Press, New York.
Bushnell, J. H., Rao, K. S. 1974. Dormant or quiescent stages and
structures among the Ectoprocta: physical and chemical factors
affecting viability and germination of statoblasts. Transactions
of the American Microscopical Society 93:524 – 543.
Canning, E. U., Okamura, B., Curry, A. 1996. Development of a
myxozoan parasite Tetracapsula bryozoides gen. n. et sp. n. in
Cristatella mucedo (Bryozoa: Phylactolaemata). Folia Parasitologica 43:249 – 261.
Canning, E. U., Okamura, B., Curry, A. 1997. A new microsporidium, Nosema cristatellae n. sp. in the bryozoan Cristatella
mucedo (Bryozoa, Phylactolaemata). Journal of Invertebrate
Pathology 70:177 – 183.
Cohen, B. L., Gawthrop, A. B. 1996. Brachiopod molecular phylogeny. Pages 73 – 80, in: Cooper, P., Jin, J. Eds. Brachiopods.
Proceedings of the Third International Brachipod Congress.
Balkema, Rotterdam.
Cooper, C. M., Burris, J. W. 1984. Bryozoans — possible indicators
of environmental quality in Bear Creek, Mississippi. Journal of
Environmental Quality 13(1):127 – 130.
Curry, M. G., Everitt, B., Vidrine, M. F. 1981. Haptobenthos on
shells of living freshwater clams in Louisiana. The Wassman
Journal of Biology 39:56 – 62.
Cusak, T. M., McCullough, J. D. 1985. Urnatella gracilis (Entoprocta) from Caddo Lake, Texas and Louisiana. The Texas
Journal of Science 37:141 – 142.
Dehdashti, B., Blinn, D. W. 1986. A bryozoan from an unexplored
cave at Montezuma Well, Arizona. The Southwestern Naturalist
31:557 – 558.
Dendy, J. S., 1963. Observations on bryozoan ecology in farm ponds.
Limnology and Oceanography 8:478 – 482.
Dendy, J. S., Sublette, J. E. 1959. The Chironomidae ( Tendipedidae: Diptera) of Alabama with descriptions of six new species.
Annals of the Entomological Society of America 52:506 – 519.
Eckman, J. E., Okamura, B. 1998. A model of particle capture by
bryozoans in turbulent flow: significance of colony form. The
American Naturalist 152(6):861 – 880.
Emig, C. 1984. On the origin of the Lophophorata. Zeitschrift für
Zoologische Systematik und Evolutionforschung 22:91 – 94.
Emschermann, P. 1965. Über die dexualle Fortpflanzung und die
Larve von Urnatella gracilis Leidy (Kamptozoa). Zeitshcrift für
Morphologie und Ökologie der Tiere 55:859 – 914.
Eng, L. L. 1977. The freshwater entoproct, Urnatella gracilis Leidy,
in the Delta – Mendota Canal, California. The Wassman Journal
of Biology 35:196 – 202.
Everitt, B. C. 1975. Fresh-water Ectoprocta: distribution and ecology
of five species in southeastern Louisiana. Transactions of the
American Microscopical Society 94:130 – 134.
Franzén, A., Sensenbaugh, T. 1983. Fine structure of the apical plate
in the larva of the freshwater bryozoan Plumatella fungosa (Pallas) (Bryozoa: Phylactolaemata). Zoomorphology 102:87 – 98.
Freeland, J. R., Jones, C. S., Noble, L. R., Okamura, B. 1999. Polymorphic microsatillite loci identified in the highly clonal freshwater bryozoan, Cristatella mucedo. Molecular Ecology 9:341 – 342.
Gilmour, T. H. J. 1978. Ciliation and function of the food-collecting
and waste-rejecting organs of lophophorates. Canadian Journal
of Zoology 56:2142 – 2155.
Goethals, B., Voss-Foucart, M. F., Goffinet, G. 1984. Composition
chimique et ultrastructure de l’ectocyste et de la coque des
524
Timothy S. Wood
statoblastes de Plumatella repens et Plumatella fungosa
(bryozoaires phylactolémes). Annales des Sciences Naturelles,
Zoologie, Paris 13e Serie 6:197 – 206.
Gutmann, W. F. 1978. Brachiopods: biomechanical interdependencies
governing their origin and phylogeny. Science 100:890 – 893.
Halanych, K. M., Bacheller, J. D., Aguinaldo, A. M. A., Liva, S. M.,
Hills, D. M., Lake, J. A. 1995. Evidence from 18S ribosomal
DNA that the lophophorates are protostome animals. Science
267:1641 – 1643.
Henry, V., Bussers, J. C., Bouquegneau, J. M., Thomé, J. P. 1989.
Heavy metal and PCB contamination of bryozoan colonies in
the River Meuse (Belgium). Hydrobiologia 202:147 – 152.
Hubschman, J. H. 1970. Substrate discrimination in Pectinatella
magnifica Leidy (Bryozoa). Journal of Experimental Biology
52:603 – 607.
Hull, H. C., Bartos, L. F., Martz, R. A. 1980. Occurrence of
Urnatella gracilis Leidy in the Tampa Bypass Canal, Florida.
Florida Scientist 43:2 – 13.
Hyman, L. H. 1959. The lophophorate coelomates: Phylum Ectoprocta. The Invertebrates. McGraw – Hill, New York.
Jebram, D. 1973. The importance of different growth directions in
the Phylactolaemata and Gymnolaemata for reconstructing the
phylogeny of the Bryozoa, in: Larwood, G. P. Ed., Living and
fossil Bryozoa. Academic Press, London.
Jones, C., Okamura, B., Noble, L. R. 1994. Parent and larval RAPD
fingerprints reveal outcrossing in freshwater bryozoans. Molecular Ecology 3:193 – 199.
Jones, K., Marsh, T., Wood, T. 2000. Surveying for phylactolaemate
bryozoans by seiving lentic sites for their statoblasts, in: Herrera
Cubilla, A. and Jackson, J. B. C., Eds., Proceedings of the 11th
International Bryozoology Association Conference, Smithsonian
Tropical Research Institute, Panama, pp. 259 – 264.
Job, P. 1976. Intervention des populations de Plumatella fungosa
(Pallas) (Bryozoaire phylactoléme) dans l’autoépuration des
eaux d’unétang et d’un ruisseau. Hydrobiologica 48:257 – 261.
Kaminski, M. 1984. Food composition of three bryozoan species
(Bryozoan Phylactolaemata) in a mesotrophic lake. Polskie
Archiwum Hydrobiologii 31(1):45 – 53.
Kaminski, M. 1991a. Feeding of the freshwater bryozoan Plumatella
fungosa (Pall.). 1. Food composition and particle size selection.
Acta Hydrobiologica 33(3 / 4):229 – 239.
Kaminski, M. 1991b. Feeding of the freshwater bryozoan Plumatella
fungosa (Pall.). 2. Filtration rate, food assimilation, and production of faeces. Acta Hydrobiologica 33(3 / 4):241 – 251.
King, D. K., King, R. H., Miller, A. C. 1988. Morphology and ecology of Urnatella gracilis Leidy, (Entoprocta), a freshwater
macroinvertebrate from artificial riffles of the Tombigbee River,
Mississippi. Journal of Freshwater Ecology 4(3):351 – 359.
Mackey, L. Y., Winnepenninckx, B., De Wachter, R., Backeljau, T.,
Emschermann, P., Garey, J. R. 1996. 18S rRNA suggests that
Entoprocta are protostomes, unrelated to Ectoprocta. Journal
of Molecular Evolution 42:552 – 559.
Marcus, E. 1926. Beobachtungen und Versuche an lebenden
Süßwasserbryozoen. Zoologische Jahrbücher. Abteilung für Systematik, Ökologie und Geographie der Tiere 52:279 – 350.
Marcus, E. 1934. Uber Lophopus crystallinus (Pall.). Zoologische
Jahrbücher. Abteilung für Anatomie und Ontogenie der Tiere
58:501 – 606.
Marcus, E. 1941. Sôbre Bryozoa do Brasil. Boletim da Faculdade de
Filosopia, Ciências e Letras, Universidade de São Paulo
5:3 – 208.
Massard, J., Geimer, G. 1996. On the occurrence of Fredericella indica Annandale, 1909 (Phylactolaemata) in Europe, in: Gordon,
D., Smith, A., Grant-Mackie, J. Eds., Bryozoans in space and
time. National Institute of Water & Atmospheric Research,
Wellington, New Zealand, pp. 187 – 192.
Massard, J. A., Geimer, G., Bromley, H. J., and Dimentman, C.
1992. Additional note on the fresh and brackish water Bryozoa
of Israel (Phylactolaemata, Gymnolaemata). Bull. Soc. Nat. luxemb. 93: 199 – 214.
Meacham, R. H., Jr., Woolcott, W. S. 1968. Studies of the coelomic
fluid and isotonic homogenates of the freshwater bryozoan
Lophopodella carteri (Hyatt) on fish tissues. Virginia Journal of
Science 19(3):143 – 146.
Morse, W. 1930. The chemical constitution of Pectinatella. Science
71:265.
Mukai, H. 1982. Development of freshwater bryozoans (Phylactolaemata), in: Harrison, F. W., Cowden, R. R. Eds. Developmental
biology of freshwater invertebrates. Alan R. Liss, New York,
pp. 535 – 576.
Mukai, H., Oda, S. 1980. Histological and histochemical studies on
the epidermal system of higher phylactolaemate bryozoans. Annotationes Zoologicae Japonenses 53:1 – 17.
Mukai, H., Tsuchiya, M., Kimoto, K. 1984. Fusion of ancestrulae
germinated from statoblasts in plumatellid freshwater bryozoans. Journal of Morphology 179:197 – 202.
Mukai, H., Fukushima, M., Jinbo, Y. 1987. Characterization of the
form and growth pattern of colonies in several freshwater bryozoans. Journal of Morphology 192:161 – 179.
Mukai, H., Kobayashi, K. 1988. External observations on the formation of statoblasts in Plumatella emarginata (Bryozoa, Phylactolaemata). Journal of Morphology 196:205 – 216.
Neck, R., Fullington, R. 1983. New records of the freshwatyer ectoproct Pectinatella magnifica in eastern Texas. The Texas Journal
of Science 35(3):269 – 271.
Nielsen, C. 1977. Phylogenetic considerations: the protostomian
relationships, in: Woollacott R., Zimmer, R. Eds., Biology of
Bryozoans. Academic Press, New York.
Oda, A. 1980. Effects of light on the germination of statoblasts in freshwater Bryozoa. Annotationes Zoologicae Japonenses 53:238–253.
Oda, S. 1982. Urnatella gracilis, a freshwater kamptozoan occurring
in Japan. Annotationes Zoologicae Japonenses 55(3):151 – 166.
Okamura, B. 1996. Occurrence, prevalence, and effects of the
myxozoan Tetracapsula bryozoides parasitic in the freshwater
bryozoan Cristatella mucedo (Bryozoa: Phylactolaemata). Folia
Parasitologica 43:262 – 266.
Okamura, B. 1997. The ecology of subdivided populations of a
clonal freshwater bryozoan in southern England. Archiv für
Hydrobiologie 141(1):13 – 34.
Okamura, B., Doolan, L. A. 1993. Patterns of suspension feeding in
the freshwater bryozoan Plumatella repens. Biological Bulletin
184:52 – 56.
Okamura, B., C., Jones, S., Noble, L. R. 1993. Randomly amplified
polymorphic DNA analysis of clonal population structure and
geographic variation in a freshwater bryozoan. Proceedings of
the Royal Society of London 253:147 – 154.
Protosov, A. A. 1980. On distribution of Urnatella gracilis (Kamptozoa) with reference to discharges of heated waters by
thermal electrical power stations. Zoologicheskii Zhurnal
59:1569 – 1571 (in Russian).
Rao, K. S., Bushnell, J. H. 1979. New structures in binding designs
of freshwater Ectoprocta dormant bodies (statoblasts). Acta
Zoologica (Stockholm) 60:123 – 127.
Rao, K. S., Diwan, A. P., Shrivastava, P. 1978. Structure and environmental relations of sclerotized structures in fresh water
Bryozoa. III. Observations on Plumatella casmiana (Ectoprocta:
Phylactolaemata). Journal of Animal Morphology and Physiology 25:8 – 15.
14. Bryozoans
Reynolds, J. D. 1976. Occurrence of the fresh-water Bryozoan,
Cristatella mucedo Cuvier, in British Columbia. Syesis 9: 365–366.
Ricciardi, A., Reiswig, H. 1994. Taxonomy, distribution, and ecology
of the freshwater bryozoans (Ectoprocta) of eastern Canada.
Canadian Journal of Zoology 72:339 – 359.
Ricciardi, A. and Wood, T. S. 1992. Statoblast morphology and systematics of the freshwater bryozoan Hyalinella orbisperma
(Kellicott, 1882). Can. J. Zool. 70:1536 – 1540.
Richelle, E., Moureau, Z., Van de Vyver, G. 1994. Bacterial feeding
by the freshwater bryozoan Plumatella fungosa (Pallas, 1768).
Hydrobiologia 291:193 – 199.
Rüsche, E. 1938. Nahrungsaufname und Nahrungsauswertung bei
Plumatella fungosa (Pallas). (Ein Beitrag zur Frage des Stoffkreislaufs im Weiher). Archiv für Hydrobiolgie 33:271 – 293.
Schröder, O. 1912. Zur Kenntnis der Buddenbrockia plumatellae
Ol. Schröder. Zeitschrift für wissenschafliche Zoologie 102:
79 – 91.
Shrivastava, P., Rao, K. S. 1985. Ecology of Plumatella emarginata
(Ectoprocta: Phylactolaemata) in the surface waters of Madhya
Pradesh with a note on its occurrence in the protected waterworks of Bhopal (India). Environmental Pollution (Series A)
39:123 – 130.
Silén, L. 1954. Developmental biology of Phoronidea of the Gullmar
Fiordarea (west coast of Sweden). Acta Zoologica 31:215 – 257.
Sládeček, V. 1980. Indicator value of freshwater Bryozoa. Acta Hydrochimica and Hydrobiologica 8:273 – 276.
Smith, D. G. 1989a. On Stephanella hina Oka (Ectoprocta: Phylactolaemata) in North America, with notes on its morphology and systematics. Journal of the American Benthological Society 7:253–259.
Smith, D. G. 1989b. Keys to the freshwater macroinvertebrates of
Massachusetts. No. 4: Benthic colonial phyla, including the
Cnidaria, Entoprocta, and Ectoprocta (colonial hydrozoans,
moss animals). Massachusetts Division of Water Pollution Control. Westborough, MA.
Smith, D.G. 1992. A new freshwater mos animal in the genus Plumatella (Ectoprocta: Phylactolaemata) from New England
(U.S.A.). Canadian Journal of Zoology 70:2192 – 2201.
Tenney, W. R., Woolcott, W. S. 1966. The occurrence and ecology of
freshwater bryozoans in the headwaters of the Tennessee, Savannah, and Saluda River systems. Transactions of the American Microscopical Association 85:241 – 245.
Vinogradov, A. V. 1996. New fossil freshwater bryozoans from the
Asiatic part of Russia and Kazakhstan. Paleontological Journal
30(3):284 – 292.
Walburg et al. 1971. in Hall, Gordon E. Ed., Reservoir fisheries and
limnology. Special Publication #8. American Fisheries Society,
Washington, DC, pp. 449 – 467.
525
Wayss, K. 1968. Quantitative Untersuchungen über Wachstum und
Regeneration bei Plumatella repens (L.). Zoologische Jahrbücher.
Abteilung für Anatomie und Ontogenie der Tiere 85:1 – 50.
Weise, J. G. 1961. The ecology of Urnatella gracilis Leidy: Phylum
Entoprocta. Limnology and Oceanography 6:228 – 230.
Wilde, G. A., Burke, T. A., Keenan, C. 1981. Bryozoa from Lake
Mead and Lake Mohave, Nevada-Arizona. The Southwestern
Naturalist 26:201.
Willmer, P. 1990. Invertebrate relationships. Cambridge University
Press, Cambridge.
Wood, T. S. 1973. Colony development in species of Plumatella and
Fredericella (Ectoprocta: Phylactolaemata), in: Boardman, R. S.,
Cheetham, A., Oliver, J. Eds., Development and function of animal colonies through time. Dowden, Hutchinson and Ross,
Stroudsburg, PA, pp. 395 – 432.
Wood, T. S. 1989. Ectoproct bryozoans of Ohio. Bulletin of the Ohio
Academy of Science, New Series 8:1 – 66.
Wood, T. S. 1996. Aquarium culture of freshwater invertebrates. The
American Biology Teacher 58(1):46 – 50.
Wood, T. S. and Wood, L. J. 2000. Statoblast morphology in historical specimens of freshwater bryozoans, in: Herrera Cubilla, A.
and Jackson, J. B. C., Eds., Proceedings of the 11th International Bryozoology Association Conference, Smithsonian Tropical Research Institute, Panama, pp. 421 – 430.
Wood, T. S., Marsh, T. G. 1996. Sineportella forbesi, a new victorellid bryozoan from Illinois (Ectoprocta: Ctenostomata).
Journal of the North American Benthological Society
15(4):610 – 614.
Wood, T. S., Marsh, T. G. 1999. Biofouling of wastewater treatment
plants by the freshwate bryozoan, Plumatella vaihiriae (Hastings, 1929). Water Research 33(3):609 – 614.
Wöss, E. 1996. Life history variation in freshwater bryozoans, in:
Gordon, D., Smith, A., Grant-Mackie, J. Eds., Bryozoans in
space and time. National Institute of Water & Atmospheric
Research, Wellington, New Zealand, pp. 391 – 399.
Wöss, E. 1999. Colonization and development of freshwater bryozoan communities on artificial substrtes in the Laxenburg
Pond (Lower Austria). Proceedings of the 11th International
Bryozoology Association Conference, 1999 (in press).
Zimmer, R. L. 1973. Morphological and developmental affinities of
the lophophorates. in: Larwood, G. P., Ed., Living and fossil
Bryozoa. Academic Press, London, pp. 593 – 599.
Zimmerman, M. 1979. Larval release and settlement behavior in
three species of plumatellid Bryozoa (Ectoprocta). M.S. Thesis,
Wright State University, Dayton, OH.
This Page Intentionally Left Blank
15
TARDIGRADA
Diane R. Nelson
Department of Biological Sciences
East Tennessee State University
Johnson City, Tennessee 37614
I. Introduction
II. Anatomy
A. Cuticle
B. Claws
C. Digestive System
D. Other Anatomical Systems
III. Physiology (Latent States)
IV. Reproduction and Development
A. Reproductive Modes
B. Reproductive Apparatus
C. Mating and Fertilization
D. Eggs
E. Parthenogenesis
F. Hermaphroditism
G. Development
I. INTRODUCTION
Since 1773, when J. A. E. Goeze reported the first
observation of a “Kleiner Wasser Bär,” tardigrades
have commonly been called “water bears” (Goeze,
1773). Their bearlike appearance, legs with claws, and
slow lumbering gait formed the basis for the descriptive
name. In 1776, the naturalist Lazzaro Spallanzani introduced the name “il Tardigrado” (slow-stepper) to
describe the slow, tortoiselike movement of the animal
(Spallanzani, 1776). The systematic position of the
Tardigrada, the taxon name assigned to the group by
Doyère (1840), has ranged from inclusion within Infusoria to the Annelida to the Arthropoda (Acarina,
Crustacea, Insecta). Tardigrada was acknowledged as a
phylum by Ramazzotti (1962) in his first monograph,
confirming the position of other workers (Richters,
1926). The phylogenetic affinities of the phylum, based
on morphological studies, were clarified by recent molecular studies with 18S rRNA that showed tardigrades
to be a sister group of the arthropods (Garey et al.,
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Text Copyright © 2001 by Academic Press. Figures Copyright © 2001 by Diane R. Nelson
All rights of reproduction in any form reserved.
V. Ecology
A. Molting, Life History, and
Cyclomorphosis
B. Habitats
C. Population Density
D. Distribution
E. Dissemination
VI. Techniques for Collection, Extraction,
and Microscopy
VII. Identification
A. Taxonomic Key to Genera of
Freshwater and Terrestrial
Tardigrada
Literature Cited
1996; Giribet et al., 1996; Garey et al., 1999), although
one study was inconclusive (Moon and Kim, 1996).
The Ecdysozoa was recently established as a new clade
containing the molting animals: arthropods, tardigrades, onychophorans, nematodes, nematomorphs, kinorhynchs, and priapulids (Aguinaldo et al., 1997).
This phylogenetic analysis, which showed no support
for the Articulata (a clade of segmented animals uniting
annelids and arthropods), offers an explanation for
morphological similarities of tardigrades to both the
onychophoran-arthropod and the aschelminth complex.
Discussion on phylogenetic relationships of tardigrades
and arthropods has been reviewed by Nielsen (1995)
and Dewel and Dewel (1997). Thus far, about 800
species of tardigrades have been described from marine,
freshwater, and terrestrial habitats; more species are expected as new areas are investigated (McInnes, 1994;
Kinchin, 1994; McInnes and Pugh, 1998).
Tardigrades are hydrophilous micrometazoans
with a bilaterally symmetrical body and four pairs of
lobopodous legs usually terminating in claws (Figs. 1
527
528
Diane R. Nelson
FIGURE 2
Scanning electron micrographs of a eutardigrade.
Macrobiotus tonollii. [From Nelson, 1982a.]
FIGURE 1
Scanning electron micrographs of a heterotardigrade.
Echiniscus spiniger. a, Cirrus A; b, buccal cirrus; c, clava; p, cephalic
papilla. [From Nelson, 1982a.]
and 2). Generally convex on the dorsal side and
flattened on the ventral side, the body is indistinctly divided into a head (cephalic) segment, three trunk segments each bearing a pair of legs, and a caudal segment
with the fourth pair of legs directed posteriorly. Although tardigrades range in body length from 50 m in
juveniles to 1200 m in adults (both excluding the last
pair of legs), mature adults average 250 – 500 m with
a few species exceeding 500 m.
Despite their overall abundance and cosmopolitan
distribution, the Tardigrada have been relatively
neglected by invertebrate zoologists. Frequently categorized as one of the “minor phyla,” they are more
appropriately termed one of the “lesser-known phyla.”
Because of difficulties in collecting and culturing the
organisms and their apparent lack of economic importance to humans, our knowledge of tardigrades has expanded slowly since their discovery over 225 years ago.
The lack of basic biological information has hindered
the growth of interdisciplinary studies, which are currently being developed.
Today, new insights into the biology of the
Tardigrada are evolving from more comprehensive, contemporary investigations utilizing molecular techniques
as well as transmission and scanning electron microscopy, phase- and interference-contrast microscopy,
and histochemical and cytological techniques. Tardigrades provide a vast array of opportunities for classical
and contemporary biological investigations. Within the
last decade, interest and activity in tardigrade research
has intensified.
II. ANATOMY
A. Cuticle
The cuticle, which is highly permeable to water,
covers the body surface and lines the foregut and
hindgut. [See Dewel et al., 1993 for detailed review.]
15. Tardigrada
TABLE I
529
Subdivision of the Phylum Tardigrada with Habitat Classificationsa
Taxon
Habitat
I. Class Heterotardigrada
A. Order Arthrotardigrada
B. Order Echiniscoidea
1. Family Echiniscoididae
2. Family Oreellidae
3. Family Carphanidae
4. Family Echiniscidae
II. Class Mesotardigrada
A. Order Thermozodia
1. Family Thermozodiidae
III. Class Eutardigrada
A. Order Parachela
1. Family Macrobiotidae
2. Family Calohypsibiidae
3. Family Eohypsibiidae
4. Family Hypsibiidae
5. Family Necopinatidae
6. Incertae sedis
B. Order Apochela
1. Family Milnesiidae
Marine, except Styraconyx hallasi (freshwater)
Marine, freshwater, terrestrial
Marine; genera Echiniscoides and Anisonyches
Terrestrial; genus Oreella
Freshwater; genus Carphania
Terrestrial, except for a few species of
Echiniscus, Hypechiniscus, and Pseudechiniscus occasionally in freshwater;
12 genera
Hot spring; genus Thermozodium
Terrestrial and freshwater; genera Adorybiotus, Calcarobiotus, Dactylobiotus, Macrobiotus, Macroversum, Microbiotus, Minibiotus, Murrayon, Pseudodiphascon, Pseudohexapodibius, Richtersius, Xerobiotus
Terrestrial; genera Calohypsibius, Hexapodibius, Haplomacrobiotus, Parhexapodibius,
Haplohexapodibius
Terrestrial and freshwater; genera Eohypsibius, Amphibolus
Marine; genus Halobiotus and a few Isohypsibius, Ramajendas. Terrestrial and freshwater;
genera Acutuneus Astatumen, Diphascon, Doryphoribius, Eremobiotus, Hebesuncus,
Hypsibius, Isohypsibius, Itaquascon, Fujiscon, Mesocrista, Microhypsibius, Mixibius,
Paradiphascon, Parascon, Platicrista, Pseudobiotus, Ramajendas, Ramazzottius,
Thulinia
Terrestrial; genus Necopinatum
Terrestrial; genus Apodibius
Terrestrial; Milnesium and Limmenius
a
Additional references for table, not cited specifically in text: Binda, 1984; Binda and Pilato, 1986; Pilato and Beasley, 1987; Pilato and
Binda, 1987a, b, 1989; Biserov, 1992; Dastych, 1992; Pilato and Binda, 1997.
Secreted by the underlying epidermis ( hypodermis),
the cuticle consists of several layers which may be further divided into: (1) an outer complex epicuticle ( exocuticle); (2) an intracuticle ( mesocuticle), which is
not present in all species; and (3) an inner procuticle (
endocuticle), which contains chitin (Greven and Peters,
1986; Greven and Greven, 1987; Dewel et al., 1993).
Although most freshwater tardigrades are white or colorless, some terrestrial species exhibit brown, yellow,
orange, pink, red, or green coloration due to intestinal
contents, body cavity cells and granules, or pigmentation in the epidermis and cuticle (Ramazzotti and
Maucci, 1983). Based on cuticular structures, three
classes, formerly considered orders, can be distinguished
(Table I). The class Heterotardigrada (Figs. 1 and 3) includes mainly the marine and armored terrestrial tardigrades, while the class Eutardigrada (Figs. 2 and 4) primarily encompasses freshwater and other terrestrial
species. A third class, Mesotardigrada (Rahm, 1937) is
based on a single species, Thermozodium esakii, from a
hot spring in Japan; it is difficult to evaluate the validity
of this class due to the lack of type specimens and to the
destruction of the type locality by an earthquake
(Ramazzotti and Maucci, 1983).
Heterotardigrades (Fig. 1) are characterized by
paired cephalic sensory appendages, which vary considerably in size and shape: internal buccal cirri, cephalic
papillae, external buccal cirri, clavae, and lateral cirri
(cirri A). Lateral cirri A mark the junction of the
cephalic and scapular plates. Some marine species also
have a median cirrus, particularly in the most primitive
order, Arthrotardigrada. In the class Eutardigrada
(Fig. 2), cephalic sensory structures, which may (Dewel
et al., 1993) or may not (Schuster et al., 1980) be homologous to those in heterotardigrades, are present
only in the order Apochela; Milnesium has two lateral
papillae and six peribuccal (oral) papillae, and Lemmenius has two lateral papillae. Other eutardigrades (order
Parachela) lack external sensory appendages but have
sensory regions on the head (Walz, 1978).
The cuticle and its processes are taxonomically important in identifying genera and species and also form
the basis for separating tardigrades into two large
groups. The “armored” tardigrades are heterotardigrades
530
Diane R. Nelson
FIGURE 3
Heterotardigrade internal anatomy (Bryodelphax
parvulus, female); lateral view (above) and ventral view (below).
[From Nelson, 1982a; drawing by R. P. Higgins.]
that have a thickened dorsal cuticle divided into individual plates, which have a species-specific pattern of sculpture. Armored tardigrades include some marine species
(families Renaudarctidae and Stygarctidae, order Arthrotardigrada) and many terrestrial species (family Echiniscidae, order Echiniscoidea). Kristensen (1987) revised the
genera of Echiniscidae, adding four new genera to the existing eight. Of these, some species in the genus Pseudechiniscus are found in freshwater, but the dorsal plates
are nonsclerotized. The “naked” or unarmored tardigrades comprise the freshwater and terrestrial eutardigrades and most marine heterotardigrades, as well as the
heterotardigrade families Oreellidae (terrestrial) and
Carphanidae (freshwater) (Binda and Kristensen, 1986;
Kristensen, 1987). Unarmored tardigrades have a thin,
smooth cuticle or a sculptured one with pores, granulation, reticulation, tubercles, papillae, or spines, but lack
plates.
B. Claws
In addition to cuticular structures, the size, shape
and number of claws (Figs. 5 and 6) are important in
FIGURE 4
Eutardigrade internal anatomy (Macrobiotus hufelandi,
female); lateral view (above) and dorsal view (below). [From Nelson,
1982a; drawing by R. P. Higgins.]
tardigrade systematics and have been used for revision
of the eutardigrades (Thulin, 1928; Marcus, 1929;
Pilato, 1969, 1982; Bertolani and Kristensen, 1987;
Kristensen and Higgins, 1984a, b; Ramazzotti and
Maucci, 1983; Bertolani and Biserov, 1996). In the heterotardigrade family Echiniscidae, each leg terminates
in four single claws in the adult and juvenile or two
claws in the larva. Frequently, the two internal claws
have a ventral spur, and occasionally one or more spurs
may be present on the external claw near the base
(Kristensen, 1987; Ramazzotti and Maucci, 1983). In
the heterotardigrade family Carphanidae, the freshwater genus Carphania has two claws on legs I – III and
only one claw on reduced leg IV; all claws are flexible
and have two basal spurs (Binda and Kristensen,
1986).
Eutardigrades usually possess two double claws on
each leg, arranged as external and internal claws
(Schuster et al., 1980; Bertolani, 1982a; Bertolani and
Pilato, 1988; Bertolani and Biserov, 1996). Each double claw consists of a basal tract and a long primary
branch with two accessory points and a secondary
branch without accessory points. In the order Apochela
15. Tardigrada
FIGURE 5 Claw types. All claws are shown in ventral view of
the fourth right leg. (A) Echiniscus; (B) Carphania; (C) Milnesium,
Limmenius; (D) Necopinatum; (E) Macrobiotus, Macroversum,
Murrayon, Pseudohexapodibius, Richtersius, Adorybiotus, Calcarobiotus, Xerobiotus; (F) Pseudodiphascon; (G) Dactylobiotus; (H)
Minibiotus; (I) Isohypsibius, Thulinia, Doryphoribius; (J) Pseudobiotus; (K) Hypsibius, Mixibius, Platicrista, Mesocrista, Hebesuncus,
Diphascon, Itaquascon, Astatumen, Fujiscon, Paradiphascon, Ramajendas; (L) Ramazzottius; (M) Calohypsibius, Microhypsibius; (N)
Hexapodibius; (O) Haplomacrobiotus; (P) Eohypsibius, Amphibolus.
a, Accessory point; b, basal tract; cb, cuticular bar; e, external claw;
i, internal claw; lu, lunule; pb, primary branch; s, spur; sb, secondary
branch. [Redrawn from Nelson and Higgins, 1990.]
(family Milnesiidae), the primary branch is long, thin,
and well separated from the stout, short secondary
branch which bears hooks or spurs. In the order
Parachela, the secondary branch and the primary
branch arise from the common basal tract. The sequence of claw branches, with respect to the midline of
extended legs, in some genera is alternate (2 – 1 – 2 – 1)
(i.e., secondary – primary – secondary – primary). In
other genera, the two primary branches are next to one
another (2-1-1-2) (i.e., secondary – primary – primary –
secondary). In some eutardigrades (e.g., Macrobiotus),
a cuticular thickening called a lunule surrounds the
base of each double claw. Lunules vary in size and
shape; they may be smooth or dentate. In several genera, one or two cuticular bars may be present on the
legs, separate from the claws but located between the
double claws, just below the base of the claws, and / or
along the sides of the internal claws.
Six major claw types have been identified based on
the genera Dactylobiotus, Macrobiotus, Calohypsibius, Hypsibius, Isohypsibius, and Eohypsibius. Additional genera have been erected based on variations in
531
FIGURE 6
Scanning electron micrographs of eutardigrade claws.
(A) Pseudobiotus; (B) Hypsibius; prim, primary branch; sec,
secondary branch; (C) Itaquascon, ap, accessory point; cb, cuticular
bar; (D) Milnesium. [From Schuster et al., 1980.]
these basic claw types. Claw reduction, particularly in
the hind claws, has evolved independently in the eutardigrades, especially those dwelling in soil habitats
(Bertolani and Biserov, 1996). In Dactylobiotus-type
claws, the two large double claws on each leg are similar in size and shape, with a 2-1-1-2 sequence, lack
lunules, and are connected at the base by a cuticular
band. The secondary branch is much shorter than the
primary and arises from a very short basal tract. The
Macrobiotus-type claws are similar in size and shape,
with a distinct peduncle (a septum separating the basal
tract from the rest of the claw), and have a 2-1-1-2 sequence, but the claws usually possess lunules and are
not connected at the base. A similar genus, Calcarobiotus, has spurs on the bases of the claws (Dastych,
1993a). In the genus Murrayon, separated from Macrobiotus by Bertolani and Pilato (1988), the claws are
V-shaped and have a quadrate peduncle (Guidetti,
1998). Macroversum is separated from Macrobiotus
by having secondary branch much shorter than the
primary branch, and the lunules are connected by a
cuticular bar (Pilato and Catanzaro, 1988; Bertolani
and Pilato, 1988). The Calohypsibius-type claws are
small and similar in size and shape but exhibit the alternate (2-1-2-1) sequence; the primary branch is
rigidly attached to the secondary branch. The Hypsibius-type claws are usually dissimilar in size and
532
Diane R. Nelson
shape, with the external double claw longer and more
slender than the internal claw. The primary branch of
the external double claw is inserted on the basal tract,
which is continuous with the secondary branch, forming a flexible junction. The Isohypsibius-type claws resemble those of Hypsibius; however, they are usually
similar in size and shape. To distinguish the two types,
the secondary branch of the external claw from one of
the first three pairs of legs should be observed in profile. In Hypsibius-type claws, the secondary branch
and basal tract run in a continuous arc or curve;
whereas, in Isohypsibius-type claws, the secondary
branch forms a right angle with the basal tract. The
genus Ramajendas was separated from Hypsibius by
having very long primary branches (Pilato and Binda,
1990). The Eohypsibius-type claws are divided into
three distinct parts: the base, the secondary branch,
and the primary branch. The branches are usually
arranged in a 2-1-2-1 sequence; however, the internal
claw can rotate 180°, resulting in a 2-1-1-2 sequence.
The external and internal claws are similar in size, but
the angle between the primary and secondary branches
is about 45° on the external claw and about 80° on
the internal claw.
C. Digestive System
1. General Anatomy
The digestive system consists of a foregut, a
midgut, and a hindgut (Figs. 3 and 4). [For a detailed
review, see Dewel et al., 1993.] Both the foregut (buccal apparatus and esophagus) and the hindgut have a
cuticular lining. The hindgut is divided into an anterior
hindgut (rectum) and a true cloaca in eutardigrades;
the opening is a ventral transverse slit just anterior to
the fourth pair of legs. In heterotardigrades, however,
reproductive and digestive systems are separate, the
hindgut is a rectum, and the gonopore opens anterior
to the anus, which is a longitudinal slit between or just
anterior to the fourth pair of legs. In the heterotardigrade family Echiniscidae, first instar individuals have a
mouth but no gonopore or anus; second instars have a
mouth and anus but no gonopore; subsequent instars
have all three openings.
2. Buccal Apparatus
The buccal apparatus is a complex structure with
considerable taxonomic significance in the eutardigrades
(Pilato, 1972, 1982; Schuster et al., 1980). Basically, it
consists of a buccal tube, a muscular sucking pharynx,
and a pair of piercing stylets that can be extended
through the mouth opening (Figs. 3,4,7,8). The position
of the mouth may be terminal (as in Macrobiotus and
FIGURE 7 Buccal apparatus. (A) Heterotardigrada, Echiniscus
(ventral); (B) Eutardigrada, Macrobiotus (ventral); (C) Eutardigrada,
Macrobiotus (lateral); (D) Carphania; (E) Macrobiotus, Macroversum, Murrayon, Dactylobiotus, Calcarobiotus, Pseudohexapodibius,
Xerobiotus; (F) Adorybiotus, Richtersius; (G) Pseudodiphascon;
(H) Minibiotus; (I) Hypsibius, Isohypsibius, Ramazzottius, Ramajendas, Mixibius, Necopinatum; (J) Doryphoribius, Apodibius; (K)
Pseudobiotus; (L) Thulinia; (M) Diphascon, Platicrista, Mesocrista,
Hebesuncus, Paradiphascon; (N) Itaquascon; (O) Milnesium; (P)
Limmenius; (Q) Calohypsibius, Microhypsibius; (R) Hexapodibius,
Haplomacrobiotus, (S) Eohypsibius; and (T) Amphibolus. a, Apophysis; bc, buccal cavity; bl, buccal lamellae; br, buccal ring; bt, buccal
tube; cp, cephalic papilla; d, droplike thickening; f, furca; ma,
macroplacoid; mic, microplacoid; p, placoid; pb, pharyngeal bulb;
pp, peribuccal papilla; s, septulum; sp, stylet support; ss, stylet
sheath, st, stylet; and vl, ventral lamina. [Redrawn from Nelson and
Higgins, 1990.]
Milnesium) or subterminal (as in Hypsibius). In some
eutardigrades, cuticular structures surround the mouth
such as elongate peribuccal papillae (6 in Milnesium),
flattened peribuccal papulae (10 in Minibiotus and Haplomacrobiotus and 6 in Calohypsibius), or lobes (6 in
Isohypsibius and others) (Schuster et al., 1980).
The buccal tube begins with a buccal ring, which
bears lamellae in some genera. The number of lamellae
varies with the genus; e.g., there are 10 in Macrobiotus,
12 in Thulinia, 14 in Amphibolus, and about 30 in
Pseudobiotus. In Milnesium, the mouth is surrounded
15. Tardigrada
533
FIGURE 8 Scanning electron micrographs of eutardigrade buccal openings. (A) Dactylobiotus,
mu, buccal mucrones; bl, buccal lamella; (B) Minibiotus, buccal opening and papulae; (C) Haplomacrobiotus, pbpu, peribuccal papulae; (D) Isohypsibius, pbl, peribuccal lobe; (E) Pseudobiotus,
buccal opening and buccal lamellae; and (F) Milnesium, cp, cephalic (lateral) papilla; pbbl, peribuccal (oral) papilla. [From Schuster et al., 1980.]
by six triangular lamellae, which close the opening.
The interior of the buccal cavity may have anterior and
posterior rows of small cuticular teeth (mucrones)
followed by dorsal and ventral transverse ridges (also
called crests or baffles), which have significant
taxonomic value as species-specific characteristics
(Pilato, 1972; Schuster et al., 1980). Most eutardi-
grades have a completely rigid buccal tube. However,
in some genera, (the Diphascon assemblage, Pseudodiphascon, Itaquascon), there is a rigid anterior buccal
tube between the buccal opening and the stylet supports, and a flexible, spiral-walled posterior pharyngeal
tube from just below the stylet supports to the pharynx. In a similar genus, Astatumen, the rigid portion of
534
Diane R. Nelson
the buccal tube is only as long as the anterior apophyses, the pharyngeal tube is spiraled, and stylet supports
are lacking (Pilato, 1997). In Macrobiotus and other
genera, there is a median ventral lamina ( buccal tube
support, reinforcement rod) that extends from the
buccal ring to the midregion of the tube.
Four major types of buccal apparatus have been described based on the genera Macrobiotus, Hypsibius,
Diphascon, and Pseudodiphascon. The Macrobiotustype has a rigid buccal tube with a ventral lamina (reinforcement rod), whereas the Hypsibius-type has a rigid
buccal tube but lacks a ventral lamina. The Diphascontype has a rigid anterior buccal tube and a flexible, annulated posterior pharyngeal tube; it also lacks a ventral
lamina. The Pseudodiphascon-type has a buccal tube
like Diphascon, but also has a ventral lamina. More
types may be identified in the future as additional genera
are described (Pilato and Binda, 1990, 1996). Although
the buccal apparatus was used by Schuster et al. (1980)
to revise the families of eutardigrades, Ramazzotti and
Maucci (1983) and subsequent authors have basically
followed Pilato’s classification based on claw structure.
The stylet mechanism consists of two protrusible
stylets, stylet sheaths, stylet supports, and associated
muscles (Schuster et al., 1980). The stylets are paired,
piercing structures lateral to the buccal tube. Composed of cuticular and calcified substances that dissolve
in some mounting media, the stylets lie in the lumen of
the salivary glands, which are dorsolateral to the pharynx. The salivary epithelium secretes the new buccal
tube, stylets, and stylet supports after the tardigrade
molts. The stylets and salivary secretions enter the anterior end of the buccal tube through stylet sheaths on
each side. Protractor and retractor muscles operate
each stylet and are attached to the furca (condyle), the
thickened base at the posterior end of each stylet. Protractor muscles extend from the furca to the buccal
tube; retractor muscles insert on the pharynx.
Some genera have anterior apophyses for the insertion of the stylet protractor muscles on the buccal tube.
In Hypsibius, the dorsal and ventral apophyses are
hook-shaped; in Isohypsibius and other genera, they
form dorsal and ventral crests. Pilato (1987) divided the
genus Diphascon into four genera based on differences
in the shape of the apophyses and other characteristics
of the buccal – pharyngeal apparatus. The stylet support
is a flexible lateral extension that attaches the furca of
the stylet to the buccal tube, but it is poorly developed in
the genus Itaquascon and absent in Astatumen (Pilato,
1997). In the heterotardigrades, the long, thin stylets
may have delicate stylet supports or may be attached to
the pharynx by a muscular band. In general, the buccal
apparatus in heterotardigrades is of less taxonomic
importance than in eutardigrades (Kristensen, 1987).
The buccal tube enters the muscular, tripartite pharyngeal bulb ( sucking pharynx, myoepithelial
bulb) (Eibye-Jacobsen, 1996), which, in most eutardigrades, is lined with three double rows of cuticular
thickenings called placoids. The placoids alternate in
position with three equal cuticular apophyses, which
are posterior to the end of the buccal tube. The larger,
anterior placoids, called macroplacoids, are present in
either two or three transverse rows. Smaller, posterior
placoids are termed microplacoids; when present, they
are arranged in a single row. In some species of the
genus Diphascon, a medial septulum is present posterior to and alternating in position with the placoids, in
the same plane of focus as the buccal tube apophyses.
The number, size, and shape of the placoids are important species characters in the eutardigrades; however,
placoids are reduced or absent in some genera (e.g.,
Itaquascon, Astatumen, Milnesium, and Limmenius).
3. Esophagus, Midgut, and Hindgut
The pharyngeal bulb is followed by a short, tripartite esophagus, which is also lined with cuticle. A mucoid secretory substance is produced by esophageal
cells, but its function is unknown.
The midgut, the largest section of the digestive
tract, is derived from endoderm and is not lined with
cuticle. There are significant differences between eutardigrades and echiniscids in the ultrastructure of the
midgut epithelium (Dewel et al., 1993). Eutardigrades
have a straight midgut, and three excretory glands
empty into the midgut – hindgut junction; in contrast,
heterotardigrades have no excretory glands at the junction of the midgut and hindgut. Food is digested in the
midgut, which has a convoluted single layer of large
epithelial cells joined by deep continuous junctions.
The intestinal content varies in color depending on the
type of food consumed, the state of the digestive
process, and the nutritional status of the animal.
The hindgut, which is lined with cuticle, receives
the contents of the midgut and is specialized for osmoregulation. The anterior hindgut or rectum probably
functions in modification of the urine from the excretory glands ( Malpighian tubules) as well as material
from the midgut. In eutardigrades, the genital ducts
empty into the posterior hindgut, which is therefore a
true cloaca. In heterotardigrades, the anus and gonopore are separate openings.
D. Other Anatomical Systems
Reviews of the morphology of other tardigrade systems are found in Greven (1980), Bertolani (1982a), Nelson (1982a), Ramazzotti and Maucci (1983), Nelson and
Higgins (1990), Dewel et al. (1993), and Kinchin (1994).
15. Tardigrada
The nervous system consists of a dorsal lobed
brain ( cerebral ganglion, supraesophageal mass),
a circumesophageal connective around the buccal tube
( ring neuropil), a “subesophageal ganglion”, a circumpharyngeal connective around the pharynx ( posterior neuropil), and four ventral ganglia joined by
paired nerve tracts (Figs. 3 and 4) (Dewel and Dewel,
1996, 1997; Wiederhöft and Greven, 1996). Many eutardigrades and some heterotardigrades have a pair of
eye spots, cup-shaped structures with pigment granules,
embedded in the dorsolateral lobes of the brain (Dewel
et al., 1993). Walz (1978) described four sensory areas
that function as chemoreceptors or mechanoreceptors
in the anterior head region of the eutardigrade Macrobiotus: the anteriolateral sensory field, the circumoral
field, the suboral sensory region, and the pharyngeal
organ. The cephalic appendages and buccal ventroposterior plates (third pair of clavae) in heterotardigrades
also function as sense organs (Kristensen and Higgins,
1984a, 1989; Dewel et al. 1993).
Tardigrade musculature has been studied in detail
in only a few species. [See Dewel et al., 1993 for a detailed review.] The muscular system consists of dorsal
and ventral longitudinal fibers, transverse muscles for
movement of the legs, stylet muscles, pharyngeal and
intestinal muscles, and muscles for defecation and deposition of eggs. Muscles are attached to the cuticle by
protrusions (apodemes) that are reformed at each
molt. The somatic muscles are single elongate cells
that insert on the body wall. Those of eutardigrades
are intermediate between smooth and obliquely striated muscles (Walz, 1974, 1975) or cross-striated in
the pharyngeal bulb (Eibye-Jacobsen, 1996), whereas
those of marine arthrotardigrades are cross-striated
(Kristensen, 1978).
According to Ramazzotti and Maucci (1983), excretion in tardigrades probably occurs in four ways: (1)
through the salivary glands at molting; (2) through
shedding the cuticle containing accumulated excretory
granules; (3) through the wall of the midgut; and (4)
through excretory glands. Eutardigrades possess three
excretory glands ( Malpighian tubules); one dorsal
and two lateral glands empty into the junction between
the midgut and hindgut. These function as excretory
and / or osmoregulatory structures in addition to the
hindgut (rectum). The glands of the marine eutardigrade Halobiotus crispae are unusually large, reflecting
its secondary adaptation from freshwater to seawater,
where it inhabits the subtidal zone and is subject to
marked changes in salinity. In echiniscids, excretory
and / or secretory organs lie above the ventro-medial
body wall at the level of the second and third pair of
legs (Dewel et al., 1992). Each is comprised of one medial and two lateral cells enclosing a pair of convoluted
535
ducts. They probably transport material in the body
cavity fluid to the cuticle.
Associated with miniaturization, specialized respiratory and circulatory systems are absent. Respiration
occurs through the cuticle. Circulation is accomplished
by the movement of fluid and coelomocytes in the body
cavity (hemocoel) when the animal moves.
III. PHYSIOLOGY (LATENT STATES)
All tardigrades are aquatic, regardless of their
specific habitat, since they require a film of water surrounding the body to be active. They are capable, however, of entering a latent state (cryptobiosis) when environmental conditions are unfavorable. Crowe (1975)
identified five types of latency: encystment, anoxybiosis, cryobiosis, osmobiosis, and anhydrobiosis. When a
tardigrade is in a latent state, metabolism, growth, reproduction, and senescence are reduced or cease temporarily, and resistance to environmental extremes such
as cold, heat, drought, chemicals, and ionizing radiation is increased. Latent states thus have a significant
impact on the ecological role of the organism. (See review by Wright et al., 1992; Kinchin, 1994.)
Encystment commonly occurs in freshwater tardigrades living in permanent pools, although cysts can also
be formed by tardigrades inhabiting soil and moss. The
conditions for encystment are not fully known; however,
We˛glarska (1957) determined that a gradual worsening
of the environment is one inducing factor for encystment
in the freshwater tardigrade Dactylobiotus dispar. Specimens encysted regularly within 12 h if placed in water
containing cyanophytes or decomposing leaves, but a
“swift worsening” of the environment caused the animals to go into an asphyxial state (anoxybiosis).
Before encystment, the tardigrade ingests large
amounts of food for storage in body cavity cells (coelomocytes). Proceeding through stages similar to molting,
the animal first ejects the buccal apparatus and enters
the simplex stage. After this process, the gut is emptied
by defecation, the hindgut lining is shed, and the organism contracts within the cuticle, becoming immobile and barrel-shaped. A cyst membrane is formed and
gradually becomes smooth and dark, replacing the old
cuticle which sloughs off. The cyst is encased in strong
walls, often opaque and encrusted with debris. Metabolism is very reduced in the cysts. Before emergence
from the cyst when environmental conditions improve,
a new cuticle is produced, including buccal apparatus
and claws. Encystment is not obligatory in the life cycle
of a tardigrade, and reasons for emergence are not fully
known. The cysts are not able to withstand high
temperatures, as in anhydrobiosis, because of the high
536
Diane R. Nelson
water content; however, the cysts can survive for over a
year in nature without totally depleting food reserves.
Anoxybiosis, or asphyxia, is a cryptobiotic state induced by low oxygen tensions in the environmental
water. The tardigrade becomes immobile, transparent,
rigid, and extended due to water absorption resulting
from loss of osmotic control. The asphyxial state is
temperature dependent; the time required for inducement varies from a few hours to several days. Some
species can survive up to 5 days in anoxybiosis; however, strictly aquatic species live only from a few hours
to 3 days in the asphyxial state. Prior to this time limit,
specimens can be revived by the addition of oxygen to
the water; return to the active state requires a few minutes to several hours. The metabolic rate of tardigrades
in anoxybiosis is unknown.
Induced by low temperatures, cryobiosis is a type
of cryptobiosis that enables tardigrades to survive
freezing and thawing, allowing limno-terrestrial tardigrades to be common in polar regions (Kristensen,
1982; De Smet and Van Rompu, 1994; Sømme, 1996).
Slow cooling rates and the formation of barrel-shaped
tuns enhance survival, and membrane protectants such
as trehalose stabilize membrane structure (Rudolph
and Crowe, 1985). [See Westh and Kristensen, 1992;
Westh et al., 1991; Ramløv and Westh, 1992, for detailed information.]
Osmobiosis is a form of cryptobiosis that results
from elevated osmotic pressures and decreased water
activity at the surface of the animal. Some species can
tolerate variations in salinity, especially intertidal marine species and euryhaline limno-terrestrial species.
However, placed in salt solutions of various ionic
strengths, most freshwater and terrestrial tardigrades
form contracted tuns (Wright et al., 1992). The marine
tardigrade Echiniscoides sigismundi becomes turgid in
freshwater but can survive up to 3 days. Activity resumes when the animals are returned to water with the
former osmotic concentrations.
Anhydrobiosis (formerly called anabiosis) is a type
of cryptobiosis induced by evaporative water loss from
terrestrial eutardigrades and echiniscids. [See Wright,
1989a, b; Wright et al., 1992; Crowe et al., 1992;
Kinchin, 1994; Sømme, 1996 for reviews.] Nearly all
marine and true freshwater tardigrades lack the ability
to survive dehydration and undergo cryptobiosis, but
experimental evidence is sparse. As the environmental
water surrounding the animal evaporates, the tardigrade contracts, retracting the head and legs and becoming the characteristic barrel-shaped tun. The rate of
desiccation must be slow to ensure survival and return
to active life with the addition of water. Survival of dehydration is correlated with synthesis of trehalose, a
disaccharide of glucose, during dehydration or its
degradation following rehydration. Trehalose alters the
physical properties of membrane phospholipids and
stabilizes dry membranes in anhydrobiotic organisms
(Crowe et al., 1984; Westh and Ramløv, 1991). In the
anhydrobiotic state, the tardigrade is highly resistant to
extreme environmental factors (temperature, radiation,
chemicals). Anhydrobiotic animals have remained viable on herbarium specimens for as long as 120 years.
The eggs of terrestrial tardigrades can also withstand
desiccation, but little is known about the eggs of the
distinctly aquatic species. The life span of anhydrobiotic tardigrades can be greatly extended by going in
and out of this cryptobiotic state, although frequent dehydration and rehydration results in reduced viability
due to the depletion of lipid reserves, which can be
converted to the membrane protectants glycerol and
trehalose (Westh and Ramløv, 1991). In the anhydrobiotic state, tardigrades can also withstand extraordinarily high hydrostatic pressure, up to 600 MPa, in perfluorcarbon (Seki and Toyoshima, 1998).
IV. REPRODUCTION AND DEVELOPMENT
A. Reproductive Modes
Sexual reproduction and parthenogenesis are reproductive modes exhibited in the Tardigrada (Ramazzotti
and Maucci, 1983; Nelson, 1982a; Bertolani, 1982b,
1983a, b, 1987a, 1990, 1992, 1994; Rebecchi and
Bertolani, 1988, 1994; Kinchin, 1994; Bertolani and
Rebecchi, 1999). Marine tardigrades are bisexual (gonochoristic) with sexual dimorphism evident in the gonopore structure; hermaphroditic species are very rare, and
parthenogenesis is absent (Bertolani, 1994). Nonmarine
species, especially the eutardigrades, are usually gonochoristic (dioecious) with unisexual or bisexual populations, but external sexual dimorphism is rarely observed.
Males are generally smaller than females; however, mature males may be larger than immature females in the
same population. Males may also exhibit modifications
of the claws (Rebecchi and Nelson, 1998), especially on
the first pair of legs, such as in Milnesium and the freshwater Pseudobiotus megalonyx and P. kathmanae (formerly P. augusti, in Kathman and Nelson, 1987, see revision in Nelson et al., 1999). These secondary sexual
characteristics arise during the molt preceding sexual
maturity (Bertolani, 1992; Rebecchi and Nelson, 1998).
In heterotardigrades, additional sexual dimorphism may
be exhibited in the length of the cephalic appendages, especially the clavae (Kinchin, 1994; Claxton, 1996). Meiotic maturation of the gametes occurs in amphimictic
tardigrades. In three species of gonochoristic freshwater
and terrestrial tardigrades, all of the females were
iteroparous, but males were either iteroparous or semelparous. In nonmarine tardigrades, unisexual populations
15. Tardigrada
are the most common type, and parthenogenesis is the
mode of reproduction, often associated with polyploidy
and with ameiotic maturation of the oocyte. Simultaneous hermaphroditism is sporadic but occurs in freshwater and terrestrial eutardigrades (Bertolani, 1994).
B. Reproductive Apparatus
A single (unpaired) gonad is present in both males
and females (Figs. 3 and 4). This saclike structure is dorsal to the intestine and attached to the body wall by two
anterior ligaments (small muscles, Dewel et al., 1993) in
adult eutardigrades or by a single median ligament
(basement membrane, Kristensen, 1979) in heterotardigrades (Nelson 1982a, Dewel et al., 1993). The morphology of the gonad varies with age, sex, species, and
the reproductive activity of the tardigrade. The male
tardigrade has two sperm ducts (vasa deferentia) that
open into the hindgut (cloaca); the female has only one
oviduct on either the right side or the left side of the intestine that opens into the hindgut. A small seminal receptacle (with internal epithelial spermatheca), which
opens ventrally into the hindgut next to the oviduct
opening, is present in only a few species of Macrobiotus,
Xerobiotus, Ramazzottius, and Hypsibius (Marcus,
1929; Bertolani, 1983a; Rebecchi and Bertolani, 1988;
Rebecchi, 1997). In eutardigrades, the cloacal opening is
a ventral transverse slit anterior to the fourth pair of
legs. In heterotardigrades, the preanal gonopore is
rosette-shaped in females but is a protruding, round or
oval-shaped aperture in males; only a few marine species
have a postanal gonopore (Kinchin, 1994). Paired external cuticular seminal receptacles (“annex glands”) lateral to the gonopore are present in females in many
species of marine heterotardigrades, but in only a few
eutardigrades. In some species of both heterotardigrades
and eutardigrades, males have seminal vesicles. (See
Dewel et al., 1993 and Kinchin, 1994 for a review.)
537
hufelandi, the nutcracker-shaped motile spermatozoa
with a helical nucleus, kidney-shaped midpiece, and
long tail are associated with internal fertilization; the
female has a spermatheca that contains straight, nonmotile spermatozoa with a cylindrical midpiece and a
very short tail (Rebecchi, 1997). In the freshwater eutardigrade Isohypsibius granulifer, transfer of modified
sperm occurs by copulation, and spermatozoa are
found in the ovary of the female (Wolburg-Buchholz
and Greven, 1979). In some heterotardigrades, the
spermatozoa are less specialized, the female has no
seminal receptacle, and fertilization is external (Kristensen, 1979). Rebecchi et al., (1999) reviewed ultrastructural differences in eutardigrade spermatozoa,
which are considered highly specialized. Male gametes
of eutardigrades are 25 – 100 m in length, usually
have an elongated head with a coiled piece (acrosome
or nucleus), a midpiece, and a tail with a tuft of 9 – 11
fine filaments. Heterotardigrade spermatozoa, considered primitive, are shorter (14 – 20 m), have a globose
head, lack a well-defined midpiece, and have a tapering
flagellum. In eutardigrades, spermatozoan morphology
can be correlated with sclerified structure of adults and
eggs, thus providing valuable taxonomic characters
(Guidi and Rebecchi, 1996).
Mating and fertilization have been observed in
only a few species of tardigrades (Ramazzotti, 1972;
Nelson, 1982a; Bertolani, 1987a, 1992). In some freshwater species, one or more males deposit sperm into
the cloacal opening of the old cuticle of the female
prior to, or during, a molt. External fertilization occurs
as the eggs are deposited in the old cuticle. In the eutardigrade Pseudobiotus megalonyx, diploid males
were able to locate mates and to distinguish between
diploid amphimictic individuals and triploid parthenogenetic ones (Bertolani, 1992). In heterotardigrades,
cephalic sensory structures may play a role in mating.
D. Eggs
C. Mating and Fertilization
Tardigrade spermatozoa have been described in
some species (Kristensen, 1979; Baccetti, 1987;
Bertolani, 1983b; Rebecchi and Guidi, 1991, 1995;
Guidi and Rebecchi, 1996; Rebecchi, 1997; Rebecchi
et al., 1999). Henneke (1911) first observed the
presence of two kinds of sperm in the freshwater tardigrade Pseudobiotus macronyx. The ultrastructure of
the spermatozoa of Macrobiotus hufelandi was first investigated by Baccetti et al. (1971). Other workers
described the morphology of spermatozoa and correlated morphology with a specialized mode of sperm
transfer (Wolburg-Buchholz and Greven, 1979; Rebecchi and Guidi, 1991, 1995; Guidi and Rebecchi, 1996;
Rebecchi, 1997). For example, in Xerobiotus pseudo-
Eggs are often essential for the identification of
freshwater and terrestrial species, especially in some
genera of eutardigrades (Fig. 9). Ornamented eggs,
such as those characteristic of Macrobiotus, Murrayon,
Minibiotus, Dactylobiotus, Amphibolus, Eohypsibius,
Ramazzottius, Hebesuncus, and some Hypsibius
species, as well as the heterotardigrade Oreella, are
usually deposited freely and singly or in small groups.
The ornamentation may include pores, reticulations,
and processes. The different types of egg ornamentations and their taxonomic significance have been discussed by various authors (Grigarick et al., 1973; Toftner et al., 1975; Greven, 1980; Ramazzotti and
Maucci, 1983; Biserov, 1990a, b; Bertolani and Rebecchi, 1993; Bertolani et al., 1996). In some genera,
538
Diane R. Nelson
smooth oval eggs with a thin shell are usually deposited
in the molted cuticle. These smooth eggs are typical of
the armored heterotardigrades, most eutardigrades in
the genera Pseudobiotus, Doryphoribius, Hypsibius,
Isohypsibius, Diphascon, Platicrista, Itaquascon, and
Milnesium, and only a few of the Macrobiotus species.
Some aquatic tardigrades deposit eggs inside the empty
exoskeletons of cladocerans, ostracods, or insects. The
aquatic eutardigrades Isohypsibius annulatus and two
species of Pseudobiotus carry the exuvium containing
eggs attached to the body for some time (Bertolani,
1982a; Rebecchi and Nelson, 1998). This is the only
known example of parental care among tardigrades.
The method of egg deposition is unknown for many
species, especially in marine tardigrades.
Echiniscus, Bryochoerus, Parechiniscus, and many
Pseudechiniscus species; males were rare in Bryodelphax, Testechiniscus, Cornechiniscus, and Mopsechiniscus but common (50% of the population) in
Oreella, Hypechiniscus, Proechiniscus, and Antechiniscus. In the absence of males in a tardigrade population,
parthenogenesis was assumed to be the mode of reproduction. Dastych (1987a), however, reported the presence of males in four species of the genus Echiniscus.
Claxton (1996) reported males to be common in a disproportionately large number of species (5 of the 8
species) of Echiniscus found in Australia; males and females were differentiated on the basis of gonopore
morphology, the size of the males, and the length of the
clavae.
Grigarick et al. (1975) reported two “types” of
gonopores and published the first scanning electron micrograph of a male gonopore of a heterotardigrade.
With regard to this report, Nelson (1982a) stated, “If
males were indeed present, this would be the first observation of males in any Echiniscus species . . . ”
Based on the presence of ventral plates, Dastych
(1987a) placed the species studied by Grigarick et al.
(1975) in Kristensen’s (1987) new genus Testechiniscus,
in which males are rare.
Polyploidy is often associated with parthenogenesis
and is frequently found in eutardigrades in various genera that inhabit different microhabitats (Bertolani,
1975, 1982b; Nelson, 1982a). Meiotic (automictic)
parthenogenesis occurs in diploid biotypes of the freshwater eutardigrades Dactylobiotus parthenogenesis and
Hypsibius dujardini. Ameiotic (apomictic) parthenogenesis occurs in triploid and tetraploid biotypes of
some freshwater and terrestrial eutardigrades (Macrobious, Xerobiotus, Pseudobiotus). According to Pilato
(1979), “ . . . many species in which males do occur
are actually an association of amphigonic with one or
more parthenogenetic, generally polyploid and reproductively isolated strains, often distinguishable by their
mechanism of egg deposition.” Since parthenogenesis
may be advantageous for invading new habitats, the
evolution of parthenogenesis may be correlated with
the evolution of cryptobiosis and the invasion of the
terrestrial habitat (Pilato, 1979; Bertolani, 1987a; Rebecchi and Bertolani, 1988; Bertolani et al., 1990;
Wright et al., 1992).
E. Parthenogenesis
F. Hermaphroditism
Parthenogenesis, a common mode of reproduction
in nonmarine tardigrades, is correlated with the ability
to undergo cryptobiosis (Bertolani, 1982b; 1994). According to Kristensen (1987), in echiniscid heterotardigrades, males had not been previously recorded in
Most eutardigrade populations are unisexual (thelytokus) although some are bisexual. Hermaphroditism
is rare in eutardigrades and known only in one marine
heterotardigrade (Bertolani, 1987a, 1994). In hermaphrodites, a single ovotestis with only one gonoduct is
FIGURE 9 Scanning electron micrographs of Macrobiotus eggs.
(A) Macrobiotus areolatus; (B) Macrobiotus hufelandi group; and
(C) Macrobiotus tonollii.
15. Tardigrada
present. Both mature and almost mature spermatozoa
and oocytes may be found in a single individual, indicating the possibility of self-fertilization. Cyclic maturation probably occurs, with the initial prevalence of
one type of gamete and then another. Hermaphroditic
eutardigrades include Parhexapodibius pilatoi from
grasslands, Macrobiotus joannae from moss, Amphibolus weglarskae from leaf litter, and four species of
Isohypsibius from freshwater (Bertolani and Manicardi, 1986).
G. Development
Developmental time, from deposition of the egg to
hatching, varies considerably within and between
species and with environmental conditions. A minimum of 5 to a maximum of 40 days have been required for egg development under experimental conditions (Marcus, 1928, 1929). Eggs can be found at any
time of the year in mosses, lichens, and soil; however,
few observations of eggs of freshwater tardigrades have
been reported (Ramazzotti and Maucci, 1983). Recently the embryology of the marine eutardigrade
Halobiotus crispae and the intertidal heterotardigrade
Echiniscoides sigismundi were investigated (Eibye-Jacobsen, 1996, 1996/97, 1997). Cleavage was total,
equal, and asynchronous; however, the cleavage pattern
and mode of mesoderm formation could not be definitely determined. Normal development of H. crispae
at 8 °C was estimated to be 15 – 16 days.
When embryonic development is complete, the immature tardigrade emerges from the egg by the action
of stylets or hindlegs or by an increase in hydrostatic
pressure created by pumping fluid into the gut with the
pharynx (Ramazzotti and Maucci, 1983; Eibye-Jacobsen, 1996/97, 1997). Postembryonic development occurs through several successive molts, primarily by the
growth of the individual cells rather than by cell division. Although tardigrades have been considered cell
constant, Bertolani (1982b) observed mitosis in eutardigrades.
In eutardigrades, development is direct and the
first-instar animals (juveniles) are similar to the adults
but have immature gonads and slight differences in the
claws and buccal apparatus. Bertolani et al. (1984)
concluded that heterotardigrades undergo indirect development characterized by two larval stages. The first
larval stage lacks an anus and a gonopore and has two
claws less on each leg than the adult. The second larval
stage (sometimes called a juvenile) has an anus and the
same number of claws as the adult but the gonopore is
rudimentary or lacking. The number of filamentous
and spinous cuticular appendages generally increases
with age (but decreases in Mopsechiniscus), although
539
variations may also occur (Grigarick et al., 1975;
Ramazzotti and Maucci, 1983; Nelson 1982a; Claxton
1996).
V. ECOLOGY
A. Molting, Life History, and Cyclomorphosis
Molting, which usually requires 5 – 10 days, occurs
periodically throughout the life of the tardigrade (Walz,
1982). The entire cuticular lining of the foregut, including the stylets and stylet supports, is ejected through
the expanded buccal opening. The mouth opening
closes and the animal cannot feed. This is the “simplex” stage, characterized by the absence of the buccal
apparatus. The salivary glands reform the cuticular
structures of the buccal tube, stylets, and stylet supports; the esophagus regenerates its own cuticular
lining. Concomitantly, new body cuticle, including the
hindgut lining, is synthesized by the underlying
epidermis and the new claws are produced by claw
glands in the legs (Walz, 1982). During the molt, the
old body cuticle is shed, including the claws and the
lining of the hindgut (Fig. 10). Generally, body length
increases with each molt until maximum size is attained, although lack of food can result in a decrease in
size. Although growth is more rapid during the earlier
molts, molting and growth continue even after sexual
maturity (Nelson 1982a). Defecation and oviposition
may also be associated with molting and changes in the
fluid pressure in the body cavity.
Life histories of certain tardigrade species have
been reviewed by Walz (1982), Nelson (1982a), Ramazzotti and Maucci (1983), and Kinchin (1994).
Based on frequency distributions of body length and
FIGURE 10
Scanning electron
Kathmanae undergoing a molt.
micrograph
of
Pseudobiotus
540
Diane R. Nelson
buccal length, the number of molts has been estimated
to range from 4 to 12. Problems inherent in the method
have been discussed by Baumann (1961), Morgan
(1977), Ramazzotti and Maucci (1983), and Kinchin
(1994). Considerable variation and overlapping of the
stages may occur within a species. Sexual maturity is
reached with the second or third molt, and egg production continues throughout life, although molting can
occur without egg production.
The lifespan of active tardigrades (excluding cryptobiotic periods) has been estimated to be from 3 – 30
months (Higgins, 1959; Franceschi et al., 1962/63; Pollock, 1970; Ramazzotti and Maucci, 1983). The total
lifespan, from hatching until death, may be greatly extended by periods of latent life (encystment and anhydrobiosis). Since aquatic tardigrades have little or no
ability to undergo anhydrobiosis, the period of active
life generally corresponds to that of total life. Ramazzotti and Maucci (1983) concluded that freshwater
species of Macrobiotus and Hypsibius live about 1 – 2
yr.; whereas moss-inhabiting species of the same genera
average 4 – 12 yr. Shorter lifespans of 3 – 4 months were
proposed by Pollock (1970) for marine species and
Franceschi et al. (1962/63) for moss-inhabiting species.
Morgan (1977) estimated a lifespan of 3 – 7 months for
Macrobiotus hufelandi and up to 3 months for Echiniscus testudo in roof mosses.
Cyclomorphosis, an annual cycle of morphological
change in individuals, has been documented in the marine eutardigrade Halobiotus crispae (Kristensen 1982).
A sexually mature summer morph alternates with a
sexually immature winter morph that tolerates freezing
temperatures and low salinities. Cyclomorphosis may
complicate identifications, especially since the distribution of the phenomenon is unknown.
B. Habitats
Although all active individuals require water, the
environments in which tardigrades live are generally divided into: (1) marine and brackish water; (2) freshwater; and (3) terrestrial habitats. The distinction between
freshwater and terrestrial species is sometimes unclear
because some tardigrades can live in a wide range of
habitats.
With few exceptions, marine tardigrades belong to
the class Heterotardigrada, either in the order Arthrotardigrada or the order Echiniscoidea (Table I). In the
class Eutardigrada, only five species of the genus Halobiotus (formerly Isohypsibius) and a few species of
Isohypsibius and Ramajendas are marine (Crisp and
Kristensen, 1983; Pilato and Binda, 1990). Although
relatively few studies on marine tardigrades have been
conducted, they indicate high morphological diversity,
with most of the genera having one or two species
(Renaud-Mornant, 1982, 1987; Grimaldi De Zio et al.,
1983; Villora-Moreno and De Zio Grimaldi, 1996).
Marine tardigrades inhabit the intertidal zone and
shallow waters off the continental shelf and as well as
bathyl and abyssal depths, including manganese
nodules in the deep sea of the eastern South Pacific
(Bussau, 1992). Three major ecological groups of
marine tardigrades are recognized:
1. a small number of species on intertidal algae or
other substrates (e.g., Echiniscoides on barnacles),
including facultative and obligatory invertebrate
ectoparasites (Tetrakentron on a sea cucumber)
and commensals (e.g., Actinarctus, Pleocola);
2. a larger group of species inhabiting the interstitial biotype, particularly in the top few
centimeters of sand near the low tide mark
(e.g., Batillipes); and
3. a highly diversified group of abyssal and deepsea ooze dwellers (e.g., Coronarctus).
Hydrophilous tardigrades are “distinctly aquatic”
species that live only in permanent freshwater habitats.
Although they are occasionally found in plankton samples, they are benthic organisms that crawl on the surfaces of aquatic plants or in the interstitial spaces of
sandy sediments in ponds, lakes, rivers, and streams
(Schuster et al., 1977; Wainberg and Hummon, 1981;
Kathman and Nelson, 1987; Nelson et al., 1987; Van
Rompu and De Smet, 1988, 1991; Van Rompu et al.,
1991a, b, 1992; Strayer et al., 1994). Most species live
in the littoral zone; however, some individuals have
been collected from lakes up to 150 m deep (Ramazzotti and Maucci, 1983). Benthic algal mats in maritime Antarctic lakes and pools are productive habitats
for tardigrades (McInnes, 1995; Everitt, 1981; McInnes
and Ellis-Evans, 1987, 1990). Cryoconite holes in glaciers, formed when heat is absorbed by surface accumulation of dark dust, also provide a specialized habitat
for rotifers and tardigrades, which feed on the microflora living there (Dastych, 1993b; De Smet and Van
Rompu, 1994; Sømme, 1996). Hygrophilous tardigrades, which inhabit moist mosses, and eurytopic
species, which live in a wide range of moisture conditions, are often found in aquatic habitats.
In the Heterotardigrada, Styraconyx hallasi is the
only freshwater arthrotardigrade. In the order Echiniscoidea, Carphania fluviatilis in the family Carphanidae
(Binda and Kristensen, 1986), and a few species of
Echiniscus, Hypechiniscus, and Pseudechiniscus in the
family Echiniscidae are occasionally limnic (Kristensen,
1987). The single specimen of Echinursellus longiunguis, originally described as an arthrotardigrade from
the shores of a Chilean lake, is a cyst or cyclomorphic
15. Tardigrada
FIGURE 11
Scanning electron micrograph of an aquatic tardigrade, Pseudobiotus Kathmanae.
stage of a eutardigrade in the genus Pseudobiotus, and
is not the missing link between Arthrotardigrada and
Echiniscoidea as originally reported (Kristensen, 1987).
The status of the mesotardigrade Thermozodium esakii
from a Japanese hot spring is dubious.
The class Eutardigrada contains both terrestrial
and freshwater species (Table I). Three genera (Dactylobiotus, Pseudobiotus, and Thulinia) are exclusively
freshwater (Fig. 11). Other genera have one or more
species that are typically aquatic, i.e., Macrobiotus,
Macroversum, Mixibius, Murrayon, Amphibolus, Doryphoribius, Ramajendas, Eohypsibius, Isohypsibius,
Hypsibius, and the Diphascon group (Bertolani,
1982a; Bertolani and Pilato, 1988; Pilato and Binda,
1990; Pilato, 1992). Several species of the genus Hypsibius ( e.g., H. dujardini and H. convergens) are not obligate freshwater organisms but are often found in this
habitat. Milnesium tardigradum and Macrobiotus hufelandi are eurytopic terrestrial species that may also inhabit freshwater. The tardigrade component of the
psammon community in lakes and river banks is poorly
known, but it consists primarily of hydrophilous and
hygrophilous species that show a distinct seasonality in
population densities, with peak periods mainly in the
winter and spring (Ramazzotti and Maucci, 1983).
The largest number of tardigrade species are terrestrial and live in moist habitats, such as in soil and leaf
litter or among mosses, lichens, liverworts, and cushionshaped flowering plants (Ramazzotti and Maucci, 1983;
Kinchin, 1994). Arctic and Antarctic species also live in
terrestrial moss turfs (Dastych, 1984; Bertolani and
Kristensen, 1987; Usher and Dastych, 1987; Maucci,
1996; Miller et al., 1996). Most terrestrial tardigrades
inhabit the moss environment, which can be divided
into three groups: wet, moist, and dry mosses
(Mihelčič, 1954 / 55; Ramazzotti and Maucci, 1983).
541
Wet mosses such as submerged and emergent species in
lakes, ponds, and streams never dehydrate completely.
In contrast, moist and dry mosses living on tree trunks,
rocks, walls, roofs, and soil are subject to total desiccation for various periods of time and are dependent on
atmospheric precipitation for moisture. The typical
terrestrial habitat of a tardigrade has: (1) sufficient aeration (tardigrades are hypersensitive to low oxygen
levels); (2) alternate periods of wet and dry conditions;
and (3) suitable and sufficient food. Tardigrades are less
likely to inhabit impermeable clay soils or dense cushion mosses with thick cell walls. A distinct insular distribution of tardigrades within patches of mosses and
soil has been frequently noted (Ramazzotti and Maucci,
1983). Lichens and liverworts are inhabited by a community usually similar to that in mosses.
High species diversity and large populations of eutardigrades are also common in soil and the upper
layer of leaf litter; however, heterotardigrades are rare
or absent (Fleeger and Hummon, 1975; Manicardi and
Bertolani, 1987; Bertolani and Rebecchi, 1996;
Guidetti, 1998; Guidetti et al., in press). Interstitial eutardigrades in inorganic soils are characterized by very
short legs with small claws and a reduction or absence
of the hind claws (Bertolani and Biserov, 1996). The
tardigrade community in leaf litter from beech forests
contains some species found in freshwater, but not
those characterized by very long claws (e.g., Dactylobiotus, Pseudobiotus). There is also some overlap with
the turf community, except for turf species with very
short claws. In contrast, the litter community is significantly different from that of mosses and lichens on
beech trees in the same area (Guidetti et al., 1999.)
C. Population Density
Population densities of tardigrades are highly variable; however, neither minimal nor optimal conditions
for population growth are known. [See review in
Kinchin, 1994.] Changes in tardigrade population densities have been correlated with a variety of environmental conditions, including temperature and moisture
(Franceschi et al., 1962 / 63; Morgan, 1977; Briones et
al., 1997), air pollution (Steiner, 1994a – c, 1995), and
food availability (Hallas and Yeates, 1972). Other factors such as competition, predation, and parasitism
may also play a role. Predators include nematodes,
other tardigrades, mites, spiders, springtails, and insect
larvae; parasitic protozoa and fungi often infect tardigrade populations (Ramazzotti and Maucci, 1983).
Considerable variation in both population density and
species diversity occurs in adjacent, seemingly identical,
microhabitats. Usually 2 – 6 species are present, but 10
or more species have been found in a single sample.
542
Diane R. Nelson
In general, patchiness is a common characteristic of
tardigrade populations.
Populations of tardigrades from soil and leaf litter
habitats range from 5 to 40 individuals / cm2. [See
Nelson and Higgins, 1990 for review and Guidetti
et al., 1999.] Similarly, population densities in mosses
and other terrestrial habitats vary widely, from 0 to
50 – 200 individuals / cm2. (For review, see Ramazzotti
and Maucci, 1983; Kinchin, 1994.) Large populations
of both heterotardigrades and eutardigrades are often
found in thin, spreading mosses growing in open, exposed places; in contrast, very few heterotardigrades,
but many eutardigrades, can be collected in mosses
from moist shady areas.
Studies of the population dynamics of freshwater
tardigrades are less common than those of terrestrial
habitats (Schuster et al., 1977; Everitt, 1981; Wainberg
and Hummon, 1981; Kathman and Nelson, 1987; Nelson et al., 1987; Strayer et al., 1994). Ramazzotti and
Maucci (1983) and Nelson et al. (1987) reported that
freshwater eutardigrades often reach a very high population density in the spring, whereas Schuster et al.
(1977) and Wainberg and Hummon (1981) noted population peaks in the fall. Kathman and Nelson (1987)
stated, “Several previous life-history studies indicated
wide variation in total numbers of animals obtained
from month to month, but few studies extended beyond 12 months or presented data on replicates,
means, or standard errors for determining the validity
of these variations. Unless there are recurring patterns
over several years, abundance estimates may be invalid
if adequate numbers of quantitative replicates are not
obtained, as our data show that within-period variation was often greater than between-period variation.”
D. Distribution
Due to the paucity of data and the uncertainty of
identifications, the biogeography of the Tardigrada remains relatively unknown (McInnes, 1994). As a phylum, tardigrades have a worldwide distribution. Many
species with broad ecological requirements are considered to be cosmopolitan, whereas others with more
narrow tolerances are rare or endemic. Cluster analysis
of the distribution of extant fauna showed that limnoterrestrial tardigrades have limited generic and familial
distribution ranges defined by biogeographical
provinces and major geological events (McInnes and
Pugh, 1998; Pugh and McInnes, 1998). Distinct
Laurasian and Gondwanan familial clusters correlate
with the Triassic disintegration of Pangaea, while
separate Antarctic, Australian and New Zealand
familial / generic clusters relate to the subsequent
Jurassic / Cretaceous disintegration of Gondwana. A
“Nearctic-Arctic” biogeographic cluster is distinct from
that of “Northern and Alpine Europe”. In general,
moss-inhabiting tardigrades are more common in polar
and temperate zones than in the tropics. More data are
required, however, before definitive biogeographical distributions of the phylum can be determined.
On a more localized scale, significant differences in
altitudinal distributions of terrestrial tardigrades have
been reported by Nelson (1975), Beasley (1988),
Dastych (1987b, 1988), Manicardi and Bertolani
(1987), Ramazzotti and Maucci (1983), and Guidetti
et al. (1999). In contrast, Kathman and Cross (1991)
found that the distribution of tardigrades at any altitude is primarily determined by abiotic factors, such as
temperature and humidity, which affect the microhabitats at a particular altitude.
Physical and chemical monitoring of pollutants in
environmental surveys have been used to indicate air
and water quality; however, ecological monitoring has
not been extensively developed. Terrestrial invertebrates
in mosses and soils, as well as aquatic meiofauna, may
be suitable as a monitoring system to indicate environmental quality. Steiner (1994a – c, 1995) investigated the
influence of air pollution on moss-dwelling nematodes
and tardigrades along an urban – rural gradient near
Zürich, Switzerland. Although the abundance of tardigrades varied independently of SO2 and NO2, the number of tardigrade species decreased with increasing levels of SO2. Fumigation experiments with SO2 indicated
that the effects were dose-dependent and taxon specific.
The tardigrade Macrobiotus persimilis was typical of
sites with relatively high levels of NO2.
E. Dissemination
Eggs, cysts, and barrel-shaped anhydrobiotic specimens of terrestrial tardigrades are passively dispersed
predominantly by wind, although rain, floodwaters, and
melting snow can transport active as well as cryptobiotic
forms (Ramazzotti and Maucci, 1983). Other animals,
associated with terrestrial communities, e.g., birds,
snails, isopods, mites, centipedes, millipedes, and insects,
can also assist in the dispersal of tardigrades. Active dissemination by crawling is limited by the necessity for a
constant film of water covering the body of the tardigrade, and by its size and speed of movement.
Little information is available on dispersal of the
distinctly aquatic species, which probably undergo limited or no cryptobiosis. Transport of eggs or encysted
forms from dry temporary ponds is possible. Active
tardigrades may be distributed through river systems,
particularly during periods of heavy rainfall and flooding. The mechanism for re-population of aquatic habitats after the summer disappearance of tardigrades or
15. Tardigrada
after habitat disturbances has not been investigated
(Nelson et al., 1987).
VI. TECHNIQUES FOR COLLECTION,
EXTRACTION, AND MICROSCOPY
Qualitative collections of tardigrades from aquatic
habitats with sandy bottoms can be made by stirring
the sand in a container of water and decanting immediately after the sand settles (Schuster et al., 1977). The
decanted water is passed through a sieve (US Standard
No. 325) with pores of about 44 m, and the specimens are rinsed from the screen to a sample jar. Tardigrades in large volumes of water can be preserved by
the addition of formalin (5%) or gluteraldehyde. A
modified Boisseau apparatus has also been adapted for
tardigrade extraction (Kinchin, 1994).
Modifications of techniques to extract soil nematodes by centrifugation in water, sucrose, or Ludox®
have been used to remove tardigrades from aquatic and
terrestrial habitats (Hallas, 1975; Hallas and Yeates,
1972; Bertolani, 1982a). A watery suspension of the
sample (sediment or vegetation) is prepared and filtered
through two stacked sieves with mesh openings of
500 m or greater (top sieve) and 44 m (bottom sieve).
The sediment remaining on the fine mesh sieve is centrifuged with water at about 3000 rpm for 5 mins. The
supernatant is then poured through the fine mesh sieve,
and the remaining sediment is centrifuged with a sucrose
solution (sp. gr. 1.18) at about 5000 rpm for 1 min.
Ludox® (DuPont product, colloidal silica, Sigma –
Aldrich Chemical Co.) can be used in place of sucrose;
Ludox AM® is useful for extracting tardigrades from terrestrial / freshwater environment, whereas Ludox TM® is
more effective with marine tardigrades (McInnes, personal communication). The supernatant is immediately
rinsed through the fine mesh sieve with running tap water, and the residue is washed into a petri dish for examination under a dissecting microscope.
Collections of tardigrades from vegetation can be
made by removing the plant (moss, lichen, liverwort,
etc.) from the substrate (tree, rock, soil) and placing the
sample in a small paper bag for dry storage until processing. The sample is then placed in a container of water for several hours to reverse anhydrobiosis. The sample is agitated and squeezed to remove specimens, eggs,
and cysts. After the excess water is decanted, the remaining sediment containing the live tardigrades can be
examined with a dissecting microscope at a magnification of 30 or greater. The sample may also be washed
through sieves to remove particles smaller than 50 m
or larger than 1200 m. Boiling water or boiling
alcohol (85% or higher) added to specimens in small
543
amounts of water is adequate for tardigrade fixation,
but alcohol is recommended to preserve specimens to
be mounted later. R. D. Kathman (personal communication) found that mounting the live tardigrade directly
in Hoyer’s medium was more efficient, however longterm preservation was not as satisfactory.
Core samples of soil and leaf litter can also be
placed in paper bags for dry storage and processed according to the traditional procedure for vegetation. The
sample is thoroughly soaked in water for several hours
and examined in small quantities of thin aqueous suspensions since eggs and cysts are easily overlooked. The
technique is extremely laborious and time-consuming;
however, centrifugation with sucrose or Ludox®, as described above for aquatic samples, will reduce the time
required for extraction of tardigrades from the substrate.
After fixation, individual specimens are mounted on
microscope slides and observed with phase contrast
and / or differential interference contrast (DIC) microscopy. Hoyer’s mounting medium (distilled water,
50 ml; gum arabic, 30 g; chloral hydrate, 150 g; glycerol, 20 ml) produces cleared, distended specimens, but
the coverslip must be sealed (e.g., with epoxy paint) to
ensure permanence. The amount of chloral hydrate may
be reduced to decrease the extent of clearing of specimens, and potassium iodide (KI, 1 g) may be added to
stain specimens slightly to increase resolution of morphological structures. CMC® and CMCP® media are
useful for temporary mounts; however, excessive stain
may obscure important structures, and the specimens
may clear completely after a period of time. Other media used include Faure’s medium and polyvinyl lactophenol. More permanent mounts with balsam or synthetic resins are not as useful for phase microscopy.
Kristensen and Higgins (1984a) recommend transferring animals with a drop of 2% formalin to slides and
covering with a coverslip. The formalin preparation is
infused with a 10% solution of glycerin and 90% ethyl
alcohol and allowed to evaporate to glycerin over several days. Then the coverslip is sealed with Murrayite.
For scanning electron microscopy (SEM), fixed
tardigrades are dehydrated to absolute alcohol, transferred to an intermediary liquid, such as amyl acetate,
and dried by the critical point method (Grigarick et al.,
1975). Individuals are mounted on SEM stubs and
coated with 200 – 300 nm of gold or gold – palladium.
Tardigrades can also be purchased commercially
from Carolina Biological Supply Company (Burlington, NC) or Ward’s (Rochester, NY). Specimens from
Carolina, labeled “Milnesium” or “Milnesium and
other species” and shipped in the anhydrobiotic state,
may contain a mixture of Macrobiotus, Milnesium,
and / or Echiniscus. Cultures of aquatic specimens
shipped in the active state from Ward’s, labeled
544
Diane R. Nelson
“Hypsibius (water bear)”, were actually Thulinia in
the samples personally examined. Caution should be
exercised in relying on a company’s identifications of
specialized groups of invertebrates.
VII. IDENTIFICATION
Identification of species in the phylum Tardigrada
is based primarily on the morphology of the cuticle,
claws, buccal apparatus, and eggs, which have been
discussed in previous sections. Early European workers
described the majority of tardigrade species from temporary wet mounts of live specimens observed with the
light microscope, often resulting in inadequate, incorrect, or incomplete descriptions and illustrations. Morphology, systematics, and natural history were focal
points of their studies. Early monographs and other papers by Thulin (1928) and Marcus (1929, 1936) in
German and Cuénot (1932) in French formed the basis
of tardigrade systematics. Ramazzotti (1962, 1972)
published a monograph, in Italian, which compiled
keys and illustrated diagnoses of the species, thorough
sections on morphology and biology, and an extensive
bibliography; a supplement was produced in 1974.
The last revision of this comprehensive monograph
(Ramazzotti and Maucci, 1983) incorporated parts of
the taxonomic revisions by Pilato (1969, 1982) and
Schuster et al. (1980). An English translation of this
important monograph is available from Dr. Clark
Beasley, Department of Biology, McMurry University,
Abilene, TX. Useful references in English include those
by Morgan and King (1976), Nelson (1982a), Dewel
et al. (1993), and Kinchin (1994) and the international
symposia volumes (Higgins, 1975; We˛glarska, 1979;
Nelson, 1982b; Bertolani, 1987b; McInnes and Norman, 1996; and Greven, 1999). Kinchin (1994) reviewed the systematics, morphology, and ecology of
tardigrades and presented a guide to common species.
He synthesized classifications from various authors, dividing tardigrades artificially into “marine heterotardigrades, marine eutardigrades, limno-terrestrial heterotardigrades, and limno-terrestrial eutardigrades.”
Currently, claw structure, considered more conservative than the bucco-pharyngeal apparatus, is the major character for distinguishing genera and families
within the eutardigrades (Bertolani and Kristensen,
1987). Figures 5 – 8 illustrate the claw types and the
buccal apparatus of the major genera of eutardigrades,
basically according to the classification system of Ramazzotti and Maucci (1983), but with the inclusion of
additional genera. The figures illustrate the range of
variation in morphological structures that are considered of taxonomic importance in defining genera and
also the need for further analysis.
The following key includes common genera previously found and likely to be found in freshwater and
terrestrial habitats in North America. Positive identification with the key may be difficult because of inadequate
and often vague generic descriptions, additions of new
genera (often formed by splitting current genera), and
the need for further revision of tardigrade systematics.
A. Taxonomic Key to Common Genera of Freshwater
and Terrestrial Tardigrada
1a.
Lateral cirri A present ........................................................................................................................................................................2
1b.
Lateral cirri A absent..........................................................................................................................................................................4
2a(1a).
Cuticle without dorsal plates..............................................................................................................................................................3
2b.
Cuticle with complete or partial dorsal plates; terrestrial ...Echiniscidae [Bryochoerus, Bryodelphax, Cornechiniscus, Echiniscus, Hypechiniscus, Mopsechiniscus, Parechiniscus, Pseudechiniscus (see Kristensen, 1987 for revision).]
3a(2a).
Body indistinctly divided into eight segments; clavae present; four claws on all legs in adults; terrestrial..................................Oreella
3b.
Body eutardigrade-like in shape; clavae absent; two claws on legs I – III, one claw on legs IV; freshwater ............................Carphania
4a(1b).
Head with cephalic papillae ...............................................................................................................................................................5
4b.
Head without cephalic papillae ..........................................................................................................................................................6
5a(4a).
Head with six peribuccal papillae and two lateral papillae present; terrestrial, occasionally in freshwater...........................Milnesium
5b.
Head without peribuccal papillae; two lateral papillae present; terrestrial ..........................................................................Limmenius
6a(4b).
Claws absent on all legs; legs very short; terrestrial (Binda, 1984)........................................................................................Apodibius
6b.
Claws present on at least one pair of legs ...........................................................................................................................................7
7a(6b).
Single unbranched claws without secondary branches........................................................................................................................8
7b.
Double claws, with primary and secondary branches .........................................................................................................................9
15. Tardigrada
545
8a(7a).
One unbranched conical claw on leg I, no claws on legs II – IV; terrestrial.......................................................................Necopinatum
8b.
Two unbranched claws on legs I – III, small basal spur-like claws on leg IV; terrestrial ............................................Haplomacrobiotus
9a(7b).
Two double claws on each leg, similar in size and shape and symmetrical with respect to the median plane of the leg; claw branch
sequence is 2-1-1-2 (secondary, primary, secondary, primary) ..........................................................................................................10
9b.
Two double claws on each leg usually dissimilar in size and shape and asymmetrical with respect to the median plane of the leg;
claw branch sequence is 2-1-2-1 (secondary, primary, secondary, primary) ......................................................................................14
10a(9a).
Buccal tube of distinctly greater length between stylet supports and placoids than between stylet supports and buccal opening; posterior part of tube of spiral composition; terrestrial ..................................................................................................Pseudodiphascon
10b.
Buccal tube between stylet supports and placoids not longer than distance between stylet supports and buccal opening; tube without
spiral composition ...........................................................................................................................................................................11
11a(10b).
Stylet supports attach near middle of buccal tube; with 10 peribuccal papulae (not lamellae); terrestrial ...........................Minibiotus
11b.
Stylet supports attach in posterior half of buccal tube .....................................................................................................................12
12a(11b).
Branches of claws at right angles so that tips are remote; two claws structurally connected at base; claws without lunules; cuticle
without pores; freshwater ..............................................................................................................................................Dactylobiotus
12b.
Both branches of claw follow similar axis so that tips are close together; claws not structurally connected at base; claws
with lunules; cuticle with pores ........................................................................................................................................................13
13a(12b).
Ventral lamina (reinforcement rod, buccal tube support) on buccal tube well developed; no anterior crest on buccal tube; 10
peribuccal lamellae; terrestrial or freshwater ...................................................................................................................Macrobiotus
13b.
Ventral lamina narrow or absent; unequal dorsal and ventral crests on buccal tube; lamellae unknown; terrestrial .........Adorybiotus
14a(9b).
Buccal tube with median longitudinal anteroventral lamina (ventral lamina or buccal tube support) ..............................................15
14b.
Buccal tube without median longitudinal anteroventral lamina (ventral lamina or buccal tube support) .........................................16
15a(14a).
Legs IV reduced or absent, with no claws or one or two minute double claws; terrestrial...............................................Hexapodibius
15b.
Legs IV normally developed; terrestrial and freshwater ................................................................................................Doryphoribius
16a(14b).
Claw of Calohypsibius-type, primary and secondary branches rigidly connected, claws equal in size ..............................................17
16b.
Claw with two or three distinct parts ..............................................................................................................................................18
17a(16a).
Cuticle sculptured, two transverse rows of macroplacoids; terrestrial ............................................................................Calohypsibius
17b.
Cuticle smooth; three transverse rows of macroplacoids; terrestrial ............................................................................Microhypsibius
18a(16b).
Each double claw with three distinct parts ......................................................................................................................................19
18b.
Each double claw with two distinct parts ........................................................................................................................................20
19a(18a).
Posterior part of buccal tube of spiral composition; 14 (16?) lamellae present; terrestrial ................................................Eohypsibius
19b.
Buccal tube rigid, without spiral composition; 14 lamellae present; terrestrial or freshwater ............................................Amphibolus
20a(18b).
Buccal tube rigid, without posterior part of spiral composition .......................................................................................................21
20b.
Buccal tube with posterior part of spiral composition .....................................................................................................................25
21a(20a).
Internal and external claws of dissimilar size and shape, with secondary branch continuous with the base; no lunules; no peribuccal
lamellae . ..........................................................................................................................................................................................22
21b.
Internal and external claws of dissimilar size and shape, with secondary branch forming right angle with base; lunules
and / or peribuccal lamellae present or absent ..................................................................................................................................23
22a(21a).
Claws of oberhaeuseri-type: Hypsibius-type with internal and external claws of very dissimilar size and shape, with primary branch
of external claw very long and narrow, with very thin flexible connection to upper basal portion; elliptical dorsolateral sense organs
on head; terrestrial (Binda and Pilato, 1986) ..................................................................................................................Ramazzottius
22b.
Claws of dujardini-type: Hypsibius-type with external claw slightly larger than internal claw, with primary branch attached to median basal portion of claw; no sense organs on head; terrestrial or freshwater......................................................................Hypsibius
23a(21b).
Dorsal and ventral crests on buccal tube absent; peribuccal lunules rarely present; cuticle smooth or with granulations and / or gibbosites; terrestrial or freshwater .......................................................................................................................................Isohypsibius
23b.
Dorsal and ventral crests on buccal tube present .............................................................................................................................24
24a(23b).
With 12 peribuccal lamellae; freshwater .................................................................................................................................Thulinia
546
Diane R. Nelson
24b.
With 30 peribuccal lamellae; freshwater..........................................................................................................................Pseudobiotus
25a(20b).
Pharynx with placoids and stylet supports reduced or absent terrestrial, freshwater ...........................................................Itaquascon
25b.
Pharynx with normal placoids; buccal tube with obvious stylet supports; terrestrial or freshwater; (formerly Diphascon; revised in
Pilato, 1987) ....................................................................................................................................................................................26
26a(25b).
Apophyses for insertion of stylet muscles in shape of a “hook” .......................................................................................................27
26b.
Apophyses for insertion of stylet muscles in shape of wide, flat “ridges” .........................................................................................28
27a(26a).
Stylet apophyses in the shape of a semilunar hook; pharyngeal tube longer than the buccal tube; with (subgenus Diphascon) or
without (subgenus Adropion) droplike thickening on buccal below insertion of stylet supports..........................................Diphascon
27b.
Stylet apophyses in the shape of a blunt hook; pharyngeal tube shorter than the buccal tube ............................................Hebesuncus
28a(26b).
Stylet furcae with posteriolateral processes thickened at apices ...........................................................................................Mesocrista
28b.
Stylet furcae with posteriolateral processes spoon-like and tapering at their apices .............................................................Platicrista
LITERATURE CITED
Aguinaldo, A. M., Turbeville, J. M., Linford, L. S., Rivera, M. C.,
Garey, J. R., Raff, R., Lake, J. A. 1997. Evidence for a clade of
nematodes, arthropods, and other moulting animals. Nature
387:489 – 493.
Baccetti, B. 1987. The evolution of the sperm cell in the phylum
Tardigrada (Electron microscopy of tardigrades. 5), in:
Bertolani, R., Ed., Biology of tardigrades. Selected Symposia and
Monographs U. Z. I., 1. Mucchi, Modena, Italy, pp. 87 – 91.
Baccetti, B., Rosati, F., Selmi, G. 1971. Electron microscopy of tardigrades. 4. The spermatozoon. Monitore Zoologico Italiano
5:231 – 240.
Baumann, H. 1961. Der Lebensablauf von Hypsibius (H.) convergens
Urbanowicz (Tardigrada). Zoologischer Anzeiger 167:362 – 381.
Beasley, C. 1988. Altitudinal distribution of Tardigrada of New Mexico with the description of a new species. American Midland
Naturalist 120:436 – 440.
Bertolani, R. 1975. Cytology and systematics in Tardigrada. Memorie dell’Istituto Italiano di Idrobiologia 32 (Suppl.)17 – 35.
Bertolani, R. 1982a. Tardigradi. Guide per il riconoscimento delle
specie animali delle acque interne Italiane. Consiglio Nazionale
Delle Ricerche, Verona, Italy. 104 pp.
Bertolani, R. 1982b. Cytology and reproductive mechanisms in tardigrades, in: Nelson, D. R., Ed., Proceedings of the third international symposium on the Tardigrada. East Tennessee State University Press, Johnson City, TN, pp. 93 – 114.
Bertolani, R. 1983a. 17. Tardigrada, in: Adiyodi, K. G., Adiyodi,
R. G., Eds., Reproductive biology of invertebrates. Vol. I:
Oogenesis, oviposition, and oosorption. Wiley, New York,
pp. 431 – 441.
Bertolani, R. 1983b. 19. Tardigrada, in: Adiyodi, K. G., Adiyodi,
R. G. Eds., Reproductive biology of invertebrates. Vol. II:
Spermatogenesis and sperm function. Wiley, New York,
pp. 387 – 396.
Bertolani, R. 1987a. Sexuality, reproduction, and propagation in
tardigrades, in: Bertolani, R., Ed., Biology of tardigrades. Selected Symposia and Monographs U. Z. I., 1. Mucchi, Modena,
Italy, pp. 93 – 101.
Bertolani, R., Ed., 1987b. Biology of tardigrades. Proceedings of the
fourth international symposium on the Tardigrada, September
1985, Modena, Italy. Selected Symposia and Monographs U. Z.
I., 1. Mucchi, Modena, Italy.
Bertolani, R. 1990. Tardigrada, in: Adiyodi, K. G., Adiyodi, R. G.,
Eds., Reproductive biology of invertebrates. Vol. 4B: Fertilization,
development and parental care. Wiley, New York, pp. 49 – 60.
Bertolani, R. 1992. Tardigrada, in: Adiyodi, K. G., Adiyodi, R. G.,
Eds., Reproductive biology of invertebrates. Vol. 5B: Sexual
differentiation and behaviour. Wiley, New York, pp.
255 – 266.
Bertolani, R. 1994. 2. Tardigrada, in: Adiyodi, K. G., Adiyodi, R. G.,
Eds., Reproductive biology of invertebrates. Vol. VI: Asexual
propagation and reproductive strategies. Oxford and IBH Publ.,
New Delhi, pp. 25 – 37.
Bertolani, R., Biserov, V. 1996. Leg and claw adaptations in soil
tardigrades, with erection of two new genera of Eutardigrada,
Macrobiotidae: Pseudohexapodibius and Xerobiotus. Invertebrate Biology 115(4):299 – 304.
Bertolani, R., Kristensen, R. 1987. New records of Eohypsibius nadjae Kristensen, 1982, and revision of the taxonomic position of
two genera of Eutardigrada (Tardigrada), in: Bertolani, R., Ed.,
Biology of tardigrades. Selected Symposia and Monographs
U. Z. I., 1. Mucchi, Modena, Italy, pp. 359 – 372.
Bertolani, R., Manicardi, G. C. 1986. New cases of hermaphroditism
in tardigrades. International Journal of Invertebrate Reproduction 9:363 – 366.
Bertolani, R., Pilato, G. 1988. Struttura delle unghie nei Macrobiotidae e descrizione di Murrayon n. gen. (Eutardigrada). Animalia
15:17 – 24.
Bertolani, R., Rebecchi, L. 1993. A revision of the Macrobiotus hufelandi group (Tardigrada, Macrobiotidae), with some observations on the taxonomic characters of eutardigrades. Zoologica
Scripta 22:127 – 152.
Bertolani, R., Rebecchi, L. 1996. The tardigrades of Emilia (Italy). II.
Monte Rondinaio. A multihabitat study on a high altitude valley of the northern Apennines. Zoological Journal of the Linnean Society 116:3 – 12.
Bertolani, R., Rebecchi, L. 1999. Tardigrada, in: Knobil, E., Neill, J.
D., Eds., Encyclopedia of Reproduction, Vol. 4: Academic Press,
San Diego, pp. 703– 718.
Bertolani, R., Grimaldi De Zio, S., D’Addabbo Gallo, M., Morone
de Lucia, M. R. 1984. Postembryonic development in heterotardigrades. Monitore Zoologico Italiano 18:307 – 320.
Bertolani, R., Rebecchi, L., Beccaccioli, G. 1990. Dispersal of Ramazzottius and other tardigrades in relation to type of reproduction.
Invertebrate Reproduction and Development 18:153 – 157.
Bertolani, R., Rebecchi, L., Claxton, S. 1996. Phylogenetic significance of egg shell variation in tardigrades. Zoological Journal
of the Linnean Society 116:139 – 148.
Binda, M. G. 1984. Notizie sui Tardigradi dell’Africa Meridonale
con descrizione di una nuova specie di Apodibius (Eutardigrada). Animalia 11:5 – 15.
15. Tardigrada
Binda, M. G., Kristensen, R. 1986. Notes on the genus Oreella
(Oreellidae) and the systematic position of Carphania fluviatilis
Binda, 1978 (Carphanidae fam. nov., Heterotardigrada).
Animalia 13(1 / 3):9 – 20.
Binda, M. G., Pilato, G. 1986. Ramazzottius, nuovo genere di Eutardigrado (Hypsibiidae). Animalia 13:159 – 166.
Biserov, V. 1990a. On the revision of the genus Macrobiotus. The
subgenus Macrobiotus s.st.: a new systematic status of the
group hufelandi (Tardigrada, Macrobiotidae). Communication
1. Zoologicheskii Zhurnal 69:5 – 17 (in Russian, with English
summary).
Biserov, V. 1990b. On the revision of the genus Macrobiotus. The
subgenus Macrobiotus s.st. is a new systematic status of the
group hufelandi (Tardigrada, Macrobiotidae). Communication
2. Zoologicheskii Zhurnal 69:38 – 50 (in Russian, with English
summary).
Biserov, V. 1992. A new genus and three new species of tardigrades
(Tardigrada: Eutardigrada) from the USSR. Bolletino di Zoologia 59:95 – 103.
Briones, M. J., Ineson, P., Piearce, T. G. 1997. Effects of climate
change on soil fauna; responses of enchytraeids, Diptera larvae
and tardigrades in a transplant experiment. Applied Soil Ecology 6:117 – 134.
Bussau, C. 1992. New deep-sea Tardigrada (Arthrotardigrada,
Haleschiniscidae) from a manganese nodule area of the eastern
South Pacific. Zoologica Scripta 21:79 – 91.
Claxton, S. 1996. Sexual dimorphism in Australian Echiniscus
(Tardigrada, Echiniscidae) with descriptions of three new
species. Zoological Journal of the Linnean Society 116:13 – 33.
Crisp, M., Kristensen, R. 1983. A new marine interstitial eutardigrade from East Greenland, with comments on habitat and
ecology. Videnskabelige Meddeleser fra Dansk Naturhistorisk
Foreng 144:99 – 114.
Crowe, J. 1975. The physiology of cryptobiosis in tardigrades. Memorie dell’Istituto Italiano di Idrobiologia 32(Suppl.):37 – 59.
Crowe, J., Crowe, L., Chapman, D. 1984. Preservation of membranes in anhydrobiotic organisms: the role of trehalose. Science 223:701 – 703.
Crowe, J., Hoekstra, F., Crowe, L. 1992. Anhydrobiosis. Annual Review of Physiology 54:579 – 599.
Cuénot, L. 1932. Tardigrades, in: Lechevalier, P., Ed., Faune de
France 24:1 – 96.
Dastych, H. 1984. The Tardigrada from Antarctic with description of
several new species. Acta Zoologica Cracoviensia 27:377 – 436.
Dastych, H. 1987a. Two new species of Tardigrada from the Canadian Subarctic with some notes on sexual dimorphism in the
family Echiniscidae. Entomologische Mitteilungen aus dem Zoologischen Museum Hamburg 8 (129):319 – 334.
Dastych, H. 1987b. Altitudinal distribution of Tardigrada in Poland,
in: Bertolani, R., Ed., Biology of tardigrades. Selected Symposia
and Monographs U. Z. I., 1. Mucchi, Modena, Italy, pp.
169 – 176.
Dastych, H. 1988. The Tardigrada of Poland. Monografie Fauny Polski 16:1 – 255.
Dastych, H. 1992. Paradiphascon manningi gen. n. sp. n., a new water-bear from South Africa, with the erecting of a new subfamily
Diphasconinae (Tardigrada). Mitteilungen aus dem Hamburgischen Zoologischen Museum und Institut 89:125 – 139.
Dastych, H. 1993a. A new genus and four new species of semiterrestrial water-bears from South Africa (Tardigrada). Mitteilungen
aus dem Hamburgischen Zoologischen Museum und Institut
90:175 – 186.
Dastych, H. 1993b. Redescription of the cryoconital tardigrade
Hypsibius klebelsbergi Mihelčič, 1959, with notes on the
547
microslide collection of the late Dr. F. Mihelčič. Veröff.
Museum Ferdinandeum 73:5 – 12.
De Smet, W., Van Rompu, E. A. 1994. Rotifera and Tardigrada from
some cryoconite holes on a Spitsbergen (Svalbard) glacier. Belgian Journal of Zoology 124:27 – 37.
Dewel, R. A., Dewel, W. C. 1996. The brain of Echiniscus viridissimus
Peterfi, 1956 (Heterotardigrada): a key to understanding the phylogenetic position of tardigrades and the evolution of the arthropod head. Zoological Journal of the Linnean Society 116:35 – 49.
Dewel, R. A., Dewel, W. C. 1997. The place of tardigrades in arthropod evolution, in: Fortey, R. A., Thomas, R. H., Eds., Arthropod relationships. Systematics Association Special Volume Series 55. Chapman & Hall, London, pp. 109 – 123.
Dewel, R. A., Dewel, W. C., Roush, B. G. 1992. Unusual cuticle-associated organs in the heterotardigrade, Echiniscus viridissimus.
Journal of Morphology 212:123 – 140.
Dewel, R. A., Nelson, D. R., Dewel, W. C. 1993. Tardigrada, in:
Harrison, F. W., Rice, M. E., Eds., Microscopic anatomy of invertebrates. Vol. 12: Onychophora, Chilopoda and lesser Protostomata. Wiley – Liss, Chichester, pp. 143 – 183.
Doyère, L. 1840. Memorie sur les Tardigrades. Annales des Sciences
Naturelles, Zool., Paris, Series 2, 14:269 – 362.
Eibye-Jacobsen, J. 1996. On the nature of pharyngeal muscles cells in
the Tardigrada, In: McInnes, S. J., Norman, D. B., Eds., Tardigrade biology. Zoological Journal of the Linnean Society
116:123 – 138.
Eibye-Jacobsen, J. 1996 / 97. New observations on the embryology of
the Tardigrada. Zoologischer Anzeiger 235:201 – 216.
Eibye-Jacobsen, J. 1997. Development, ultrastructure and function
of the pharynx of Halobiotus crispae Kristensen, 1982 (Eutardigrada). Acta Zoologica 78:329 – 347.
Everitt, D. A. 1981. An ecological study of an Antarctic freshwater
pool with particular reference to Tardigrada and Rotifera. Hydrobiologia 83:225 – 237.
Fleeger, J. W., Hummon, W. D. 1975. Distribution and abundance of
soil Tardigrada in cultivated and uncultivated plots of an old
field pasture. Memorie dell’Istituto Italiano di Idrobiologia
32(Suppl.):93 – 112.
Franceschi, T., Loi, M. L., Pierantoni, R. 1962 / 63. Risultati di una
prima indagine ecologica condotta su popolazioni di Tardigradi.
Bollettino dei Musei e degli Istituti Biologici dell’Universita di
Genova 32:69 – 93.
Garey, J., Nelson, D. R., Mackey, L.Y., Li, J. 1999. Tardigrade phylogeny: congruency of morphological and molecular evidence.
Zoologischer Anzeiger 238(3 – 4): 205 – 210.
Garey, J., Krotec, M., Nelson, D. R., Brooks, J. 1996. Molecular
analysis supports a tardigrade – arthropod association. Invertebrate Biology 115:79 – 88.
Giribet, G., Carranza, S., Baguñà, J., Riutort, M., Ribera, C. 1996.
First molecular evidence for the existence of a Tardigrada
Arthropoda clade. Molecular Biology and Evolution 13:76 – 84.
Goeze, J. A. E. 1773. Uber den Kleinen Wasserbär, in: Bonnet, H. K.,
Ed., Abhandlungen aus der Insectologie, Ubers. Usw, 2.
Beobachtg, pp. 67.
Greven, H. 1980. Die Bärtierchen. Die Neue Brehm-Bucherei, Vol.
537. Ziemsen Verlag, Wittenberg Lutherstradt, Germany.
Greven, H., Ed., 1999. Proceedings of the seventh international symposium on the Tardigrada, August 1997, Dusseldorf, Germany.
Zoologischer Anzeiger 238(3 – 4): 1 – 346.
Greven, H., Greven, W. 1987. Observations on the permeability of
the tardigrade cuticle using lead as an ionic tracer, in: Bertolani,
R., Ed., Biology of tardigrades. Selected Symposia and Monographs U. Z . I., 1. Mucchi, Modena, Italy, pp. 35 – 43.
Greven, H., Peters, W. 1986. Localization of chitin in the cuticle of
548
Diane R. Nelson
Tardigrada using wheat germ agglutinin – gold conjugate as a
specific electron dense marker. Tissue and Cell 18:297 – 304.
Grigarick, A. A., Schuster, R. O., Toftner, E. C. 1973. Descriptive
morphology of eggs of some species in the Macrobiotus hufelandi group. The Pan-Pacific Entomologist 49(3):258 – 263.
Grigarick, A. A., Schuster, R. O., Toftner, E. C. 1975. Morphogenesis
of two species of Echiniscus. Memorie dell’Istituto Italiano di
Idrobiologia 32(Suppl.):133 – 151.
Grimaldi De Zio, S., D’Addabbo Gallo, M., Morone de Lucia, M. R.
1983. Marine tardigrades ecology. Oebalia 9:15 – 31.
Guidetti, R. 1998. Two new species of Macrobiotidae (Tardigrada,
Eutardigrada) from the United States of America, and some taxonomic considerations of the genus Murrayon. Proceedings of
the Biological Society of Washington 111(3):663 – 673.
Guidetti, R., Bertolani, R., Nelson, D. R. 1999. Ecological and faunistic studies on tardigrades in leaf litter of beech forests. Zoologischer Anzeiger 238(3 – 4): 215 – 223.
Guidi, A., Rebecchi, L. 1996. Spermatozoan morphology as a character for tardigrade systematics: comparisons with sclerified
parts of animals and eggs in eutardigrades, in: McInnes, S. J.,
Norman, D. B., Eds., Tardigrade biology. Zoological Journal of
the Linnean Society 116:101 – 113.
Hallas, T. E. 1975. A mechanical method for the extraction of
Tardigrada. Memorie dell’Istituto Italiano di Idrobiologia
32(Suppl.):153 – 158.
Hallas, T. E., Yeates, G. W. 1972. Tardigrada of the soil and litter of
a Danish beech forest. Pedobiologia 12:287 – 304.
Henneke, J. 1911. Beiträge zur Kenntnis der Biologie und Anatomie
der Tardigraden (Macrobiotus macronyx Duj.). Zeitschrift fuer
Wissenschaftliche Zoologie 97:721 – 752.
Higgins, R. P. 1959. Life history of Macrobiotus islandicus Richters
with notes on other tardigrades from Colorado. Transactions of
the American Microscopical Society 78:137 – 154.
Higgins, R. P., Ed., 1975. International symposium on tardigrades,
Pallanza, Italy, 17 – 19, June 1974. Memorie dell’Istituto Italiano di Idrobiologia 32(Suppl.):1 – 469.
Kathman, R. D., Cross, S. F. 1991. Ecological distribution of mossdwelling tardigrades on Vancouver Island, British Columbia,
Canada. Canadian Journal of Zoology 69:122 – 129.
Kathman, R. D., Nelson, D. R. 1987. Population trends in the
aquatic tardigrade Pseudobiotus augusti (Murray). in:
Bertolani, R., Ed., Biology of tardigrades. Selected Symposia
and Monographs U. Z. I., 1. Mucchi, Modena, Italy,
pp. 155 – 168.
Kinchin, I. M. 1994. The biology of tardigrades. Portland Press,
London, 186 p.
Kristensen, R. 1978. On the fine structure of Batillipes noerrevangi
Kristensen 1978 (Heterotardigrada). 2. The muscle-attachments
and the true cross-striated muscles. Zoologischer Anzeiger
200:173 – 184.
Kristensen, R. 1979. On the fine structure of Batillipes noerrevangi
Kristensen 1978 (Heterotardigrada). 3. Spermiogenesis, in:
Weglarska, B., Ed., Second international symposium on tardigrades, Kraków, Poland, 28 – 30, July 1977. Zeszyty Naukowe
Uniwersytetu Jagiellonskiego, Prace Zoologiczne 25:97 – 105.
Kristensen, R. 1982. The first record of cyclomorphosis in
Tardigrada based on a new genus and species from Arctic
meiobenthos. Zeitschrift für Zoologische Systematik und Evolutionsforschung 20:249 – 270.
Kristensen, R. 1987. Generic revision of the Echiniscidae (Heterotardigrada), with a discussion of the origin of the family, in:
Bertolani, R., Ed., Biology of tardigrades. Selected Symposia
and Monographs U. Z. I., 1. Mucchi, Modena, Italy, pp.
261 – 335.
Kristensen, R., Higgins, R. P. 1984a. Revision of Styraconyx
(Tardigrada: Halechiniscidae) with descriptions of two new
species from Disko Bay, West Greenland. Smithsonian Contributions to Zoology 391:1 – 40.
Kristensen, R., Higgins, R. P. 1984b. A new family of Arthrotardigrada (Tardigrada: Heterotardigrada) from the Atlantic
coast of Florida, U.S.A. Transactions of the American Microscopical Society 103:295 – 311.
Kristensen, R., Higgins, R. P. 1989. Marine Tardigrada from the
southeastern United States coastal waters. 1. Paradoxipus orzeliscoides n. gen., n. sp. (Arthrotardigrada: Halechiniscidae). Transactions of the American Microscopical Society 108: 262 – 282.
Manicardi, C., Bertolani, R. 1987. First contribution to the knowledge of Alpine grassland tardigrades. in: Bertolani, R., Ed., Biology of tardigrades. Selected Symposia and Monographs U. Z. I.,
1. Mucchi, Modena, Italy, pp. 177 – 185.
Marcus, E. 1928. Zur Embryologie der Tardigraden. Verhandlungen
der Deutschen Zoologischen Gessellschaft 32:134 – 146.
Marcus, E. 1929. Tardigrada, in: Bronn, H. G., Ed., Klassen und
Ordnungen des Tierreichs. Vol. 5. Akademische Verlagsgesellschaft, Leipzig, 608 pp.
Marcus, E. 1936. Tardigrada, in: Schultze, F., Ed., Das Tierreich.
Walter de Gruyter, Berlin, 340 pp.
Maucci, W. 1996. Tardigrada of the Arctic tundra with description
of two new species. Zoological Journal of the Linnean Society
116:185 – 204.
McInnes, S. 1994. Zoogeographic distribution of terrestrial / freshwater tardigrades from current literature. Journal of Natural History 28:257 – 352.
McInnes, S. 1995. Taxonomy and ecology of tardigrades from
Antarctic lakes. M. Phil. Thesis, Open University. 248 p.
Mclnnes, S., Ellis-Evans, J. C. 1987. Tardigrades from maritime
Antarctic freshwater lakes, in: Bertolani, R., Ed., Biology of
tardigrades. Selected Symposia and Monographs U. Z. I., 1.
Mucchi, Modena, Italy, pp. 111 – 123.
Mclnnes, S., Ellis-Evans, J. C. 1990. Micro-invertebrate community
structure within a maritime Antarctic lake. Proceedings of the
National Institute of Polar Research Symposium on Polar Biology 3:179 – 189.
McInnes, S., Norman, D., Eds., 1996. Tardigrade biology. Proceedings of the sixth international symposium on Tardigrada, August 1994, Cambridge, England. Zoological Journal of the Linnean Society 116:1 – 243.
McInnes, S., Pugh, P. 1998. Biogeography of limno-terrestrial
Tardigrada, with particular reference to the Antarctic fauna.
Journal of Biogeography 25:31 – 36.
Mihelčič, F. 1954 / 55. Zur Ökologie der Tardigraden. Zoologisher
Anzeiger 153:250 – 257.
Miller, W. R., Miller, J. D., Heatwole, H. 1996. Tardigrades of the Australian Antarctic Territories: the Windmill Islands, East Antarctica. Zoological Journal of the Linnean Society 116:175 – 184.
Moon, S. Y., Kim, W. 1996. Phylogenetic position of the Tardigrada
based on the 18S ribosomal RNA gene sequences. Zoological
Journal of the Linnean Society 116:61 – 69.
Morgan, C. 1977. Population dynamics of two species of Tardigrada,
Macrobiotus hufelandi (Schultze) and Echiniscus (Echiniscus)
testudo (Doyère), in roof moss from Swansea. Journal of Animal Ecology 46:263 – 279.
Morgan, C., King, P. 1976. British Tardigrada. Tardigrada keys and
notes for the identification of the species. Synopses of the British
Fauna, No. 9. Academic Press, London, 132 p.
Nelson, D. R. 1975. Ecological distribution of tardigrades on Roan
Mountain, Tennessee – North Carolina. Memorie dell’Istituto
Italiano di Idrobiologia 32(Suppl.):225 – 276.
15. Tardigrada
Nelson, D. R. 1982a. Developmental biology of the Tardigrada, in:
Harrison, F., Cowden, R., Eds., Developmental biology of freshwater invertebrates. Alan R. Liss, New York, pp. 363 – 368.
Nelson, D. R., Ed., 1982b. Proceedings of the third international symposium on the Tardigrada, 3 – 6, August 1980, Johnson City, TN.
East Tennessee State University Press, Johnson City, TN 236 pp.
Nelson, D. R., Higgins, R. P. 1990. Tardigrada, in: Dindal, D., Ed.,
Soil biology guide. Wiley, New York, pp. 393 – 419.
Nelson, D. R., Kincer, C. J., Williams, T. C. 1987. Effects of habitat
disturbances on aquatic tardigrade populations, in: Bertolani,
R., Ed., Biology of tardigrades. Selected Symposia and Monographs U. Z. I., 1. Mucchi, Modena, Italy, pp. 141 – 153.
Nelson, D. R., Marley, N., Bertolani, R. 1999. Re-description of the
genus Pseudobiotus (Eutardigrada, Hypsibiidae) and of the new
type species Pseudobiotus kathmanae sp. n., Zoologischer
Anzeiger 238(3 – 4): 311 – 317.
Nielsen, C. 1995. Phylum Tardigrada, in: Animal evolution — Interrelationships of the living phyla. Oxford University Press, Oxford, pp. 176 – 181.
Pilato, G. 1969. Evoluzione e nuova sistemazione degli Eutardigrada.
Bolletino di Zoologia 36:327 – 345.
Pilato, G. 1972. Structure, intraspecific variability and systematic
value of the buccal armature of eutardigrades. Zeitschrift für
Zoologische Systematik und Evolutionsforschung 10:65 – 78.
Pilato, G. 1979. Correlations between cryptobiosis and other biological characteristics in some soil animals. Bollettino di Zoologia
46:319 – 332.
Pilato, G. 1982. The systematics of Eutardigrada: A comment.
Zeitschrift für Zoologische Systematik und Evolutionsforschung
20:271 – 284.
Pilato, G. 1987. Revision of the genus Diphascon Plate, 1889, with
remarks on the subfamily Itaquasconinae (Eutardigrada, Hypsibiidae), in: Bertolani, R., Ed., Biology of tardigrades. Selected
Symposia and Monographs U. Z. I., 1. Mucchi, Modena, Italy,
pp. 337 – 357.
Pilato, G. 1992. Mixibius, nuovo genere di Hypsibiidae (Eutardigrada). Animalia 19:121 – 125.
Pilato, G. 1997. Astatumen, a new genus of the Eutardigrada (Hypsibiidae, Itaquasconinae). Entomologische Mitteilungen aus dem
Zoologischen Museum Hamburg 12:205 – 208.
Pilato, G., Beasley, C. W. 1987. Haplohexapodibius seductor n. gen.
n. sp. (Eutardigrada, Calohypsibiidae) with remarks on the systematic position of the new genus. Animalia 14:65 – 71.
Pilato, G., Binda, M. G. 1987a. Parascon schusteri n. gen. n. sp. (Eutardigrada, Hypsibiidae, Itaquasconinae). Animalia 14:91 – 97.
Pilato, G., Binda, M. G. 1987b. Richtersia, nuovo genere di Macrobiotidae, e nuova definizione di Adorybiotus Maucci & Ramazzotti 1981 (Eutardigrada). Animalia 14:147 – 152.
Pilato, G., Binda, M. G. 1989. Richtersius, nuovo nome generico in
sostituzione di Richtersia Pilato e Binda 1987 (Eutardigrada).
Animalia 16:147 – 148.
Pilato, G., Binda, M. G. 1990. Tardigradi dell’Antarctide. I. Ramajendas, nuovo genere di Eutardigrado. Nuova posizione sistematica di Hypsibius renaudi Ramazzotti, 1972 e descrizione di
Ramajendas frigidus n. sp. Animalia 17:61 – 71.
Pilato, G., Binda, M. G. 1996. Additional remarks to the description
of some genera of eutardigrades. Bollettino delle Sedute dell’Accademia Gioenia di Scienze Naturale, Catania 29:33 – 40.
Pilato, G., Binda, M. G. 1997. Acutuncus a new genus of Hypsibiidae (Eutardigrada). Entomologische Mitteilungen aus dem Zoologischen Museum Hamburg 12: 159 – 162.
Pilato, G., Catanzaro, R. 1988. Macroversum mirum n. gen. n. sp.
nuovo Eutardigrado (Macrobiotidae) dei Monti Nebrodi (Sicilia). Animalia 15:175 – 180.
549
Pollock, L. W. 1970. Distribution and dynamics of interstitial
Tardigrada at Woods Hole, Massachusetts, USA. Ophelia
7:145 – 165.
Pugh, P., McInnes, S. 1998. The origin of Arctic terrestrial and freshwater tardigrades. Polar Biology 19:177 – 182.
Rahm, G. 1937. A new ordo of tardigrades from the hot springs of
Japan (Furu-Section, Unzen). Annotationes Zoologicae Japonenses 16:345 – 352.
Ramazzotti, G. 1962. Il Phylum Tardigrada. Memorie dell’Istituto
Italiano di Idrobiologia 16:1 – 595.
Ramazzotti, G. 1972. II Phylum Tardigrada. (Seconda edizione aggiornata). Memorie dell’Istituto Italiano di Idrobiologia
28:1 – 732.
Ramazzotti, G. 1974. Supplement A, Il Phylum Tardigrada, Seconda
Edizione, 1972. Memorie dell’Istituto Italiano di Idrobiologia
31:69 – 179.
Ramazzotti, G., Maucci, W. 1983. Il Phylum Tardigrada. III edizione
riveduta e aggiornata. Memorie dell’Istituto Italiano di Idrobiologia 41:1 – 1012.
Ramløv, H., Westh, P. 1992. Survival of the cryptobiotic eutardigrade
Adorybiotus coronifer during cooling to 196°C: Effect of
cooling rate, trehalose level, and short term acclimation. Cryobiology 29:125 – 130.
Rebecchi, L. 1997. Ultrastructural study of spermiogenesis and the
testicular and spermathecal spermatozoon of the gonochoristic
tardigrade Xerobiotus pseudohufelandi (Eutardigrada, Macrobiotidae). Journal of Morphology 234:11 – 24.
Rebecchi, L., Bertolani, R. 1988. New cases of parthenogenesis and
polyploidy in the genus Ramazzottius (Tardigrada, Hypsibiidae)
and a hypothesis concerning their origin. Invertebrate Reproduction and Development 14:187 – 196.
Rebecchi, L., Bertolani, R. 1994. Maturative pattern of ovary and
testis in eutardigrades of freshwater and terrestrial habitats. Invertebrate Reproduction and Development 26:107 – 117.
Rebecchi, L., Guidi, A. 1991. First SEM studies on tardigrade spermatozoa. Invertebrate Reproduction and Development 19:151 – 156.
Rebecchi, L., Guidi, A. 1995. Spermatozoon ultrastructure in two
species of Amphibolus (Eutardigrada, Eohypsibiidae). Acta Zoologica 26(2):171 – 176.
Rebecchi, L., Nelson, D. R. 1998. Evaluation of a secondary sex character in eutardigrades. Invertebrate Biology 117(3): 194 – 198.
Rebecchi, L., Guidi, A., Bertolani, R. 1999. Tardigrada, in: Jamieson,
B. G. M., Ed., Progress in male gamete biology. vol. 9, part B.
Oxford and IBH, New Delhi.
Renaud-Mornant, J. 1982. Species diversity in marine Tardigrada, in:
Nelson, D., Ed., Proceedings of the third international symposium on the Tardigrada. East Tennessee State University Press,
Johnson City, TN., pp. 149 – 178.
Renaud-Mornant, J. 1987. Bathyl and abyssal Coronarctidae
(Tardigrada), description of new species and phylogenetical significance, in: Bertolani, R., Ed., Biology of tardigrades. Selected
Symposia and Monographs U. Z. I., 1. Mucchi, Modena, Italy,
pp. 229 – 252.
Richters, F. 1926. Tardigrada, in: Kükenthal, W., Krumbach, T., Eds.,
Handbuch de Zoologie. Walter de Gruyter, Berlin, pp. 1 – 68.
Rudolph, A. S., Crowe, J. H. 1985. Membrane stabilization during
freezing: the role of two natural cryoprotectants, trehalose and
proline. Cryobiology 22:367 – 377.
Schuster, R. O., Toftner, E. C., Grigarick, A. A. 1977. Tardigrada of
Pope Beach, Lake Tahoe, California. The Wasmann Journal of
Biology 35:115 – 136.
Schuster, R. O., Nelson, D. R., Grigarick, A. A., Christenberry, D.
1980. Systematic criteria of the Eutardigrada. Transactions of
the American Microscopical Society 99:284 – 303.
550
Diane R. Nelson
Seki, K., Toyoshima, M. 1998. Preserving tardigrades under pressure.
Nature 395:853 – 854.
Sømme, L. 1996. Anhydrobiosis and cold tolerance in tardigrades.
European Journal of Entomology 93:349 – 357.
Spallanzani, L. 1776. Opuscoli di fisica animale e vegetabile, Vol. 2,
il Tardigrado etc. Opuscolo 4:222.
Steiner, W. 1994a. The influence of air pollution on moss-dwelling
animals: 1. Methodology and composition of flora and fauna.
Revue Suisse de Zoologie 101:533 – 556.
Steiner, W. 1994b. The influence of air pollution on moss-dwelling
animals: 2. Aquatic fauna with emphasis on Nematoda and
Tardigrada. Revue Suisse de Zoologie 101:699 – 724.
Steiner, W. 1994c. The influence of air pollution on moss-dwelling
animals: 4. Seasonal and long-term fluctuations of rotifer, nematode and tardigrade populations. Revue Suisse de Zoologie
101:1017 – 1031.
Steiner, W. 1995. The influence of air pollution on moss-dwelling
animals: 5. Fumigation experiments with SO2 and exposure
experiments. Revue Suisse de Zoologie 102:13 – 40.
Strayer, D., Nelson, D. R., O’Donnell, E. B. 1994. Tardigrades from
shallow groundwaters in southeastern New York, with the first
record of Thulinia from North America. Transactions of the
American Microscopical Society 113:325 – 332.
Thulin, G. 1928. Über die Phylogenie und das System der Tardigraden. Hereditas 11:207 – 266.
Toftner, E. C., Grigarick, A. A., Schuster, R. O. 1975. Analysis of scanning electron microscope images of Macrobiotus eggs. Memorie
dell’Istituto Italiano di Idrobiologia 32(Suppl.):393 – 411.
Usher, M. B., Dastych, H. 1987. Tardigrada from the Maritime
Antarctic. British Antarctic Survey Bulletin 77:163 – 166.
Van Rompu, E. A., De Smet, W. H. 1988. Some aquatic Tardigrada
from Bjørnøya (Svalbard). Fauna Norvegica, Series A, 9:31 – 36.
Van Rompu, E. A., De Smet, W. H. 1991. Contribution to the fresh
water Tardigrada from Barentsøya, Svalbard (78°30N). Fauna
Norvegica, Series A, 12:29 – 39.
Van Rompu, E. A., De Smet, W. H., Bafort, J. M. 1991a. Some freshwater tardigrades from the Kilimanjaro. Natuurwetenschappelijk Tijdschrift 73:55 – 62.
Van Rompu, E. A., De Smet, W. H., Bafort, J. M. 1991b. Contributions to the Tardigrada of the Canadian High-Arctic. 2. Freshwater tardigrades from Little Cornwallis Island, N.W.T.
Canada. Biologisch Jaarboek Dodonaea 59:132 – 140.
Van Rompu, E. A., De Smet, W. H., Beyens, L. 1992. Contributions
to the Tardigrada of the Canadian High-Arctic 1. Freshwater
tardigrades from Devon Island, Northwest Territories. The
Canadian Field-Naturalist 106: 303 – 310.
Villora-Moreno, S., De Zio Grimaldi, S. 1996. New records of marine Tardigrada in the Mediterranean Sea. Zoological Journal of
the Linnean Society 116:149 – 166.
Wainberg, R., Hummon, W. 1981. Morphological variability of the
tardigrade Isohypsibius saltursus. Transactions of the American
Microscopical Society 100:21 – 33.
Walz, B. 1974. The fine structure of somatic muscles of Tardigrada.
Cell and Tissue Research 149:81 – 89.
Walz, B. 1975. Ultrastructure of muscle cells in Macrobiotus hufelandi.
Memorie dell’Istituto Italiano di Idrobiologia 32(Suppl.): 425–443.
Walz, B. 1978. Electron microscopic investigation of cephalic sense
organs of the tardigrade Macrobiotus hufelandi Schultze.
Zoomorphologie 89:1 – 19.
Walz, B. 1982. Molting in Tardigrada. A review including new results on cuticle formation in Macrobiotus hufelandi, in: Nelson,
D., Ed., Proceedings of the third international symposium on
the Tardigrada. East Tennessee State University Press, Johnson
City, TN, pp. 129 – 147.
We˛glarska, B. 1957. On the encystation in Tardigrada. Zoologica
Poloniae 8:315 – 325.
We˛glarska, B., Ed. 1979. Second international symposium on tardigrades, Kraków, Poland, July 28 – 30, 1977, Zeszyty Naukowe
Uniwersytetu Jagiellonskiego, Prace Zoologiczne 25:1 – 197.
Westh, P., Kristensen, R. M. 1992. Ice formation in the freezetolerant eutardigrades Adorybiotus coronifer and Amphibolus
nebulosus studied by differential scanning calorimetry. Polar
Biology 12:693 – 699.
Westh, P., Ramløv, H. 1991. Trehalose accumulation in the tardigrade Adorybiotus coronifer during anhydrobiosis. Journal of
Experimental Zoology 258:303 – 311.
Westh, P., Kristiansen, J., Hvidt, A. 1991. Ice-nucleating activity in
the freeze-tolerant tardigrade Adorybiotus coronifer. Comparative Biochemistry and Physiology [A] 73:621 – 626.
Wiederhöft, H., Greven, H. 1996. The cerebral ganglia of Milnesium
tardigradum Doyère (Apochela, Tardigrada): three-dimensional
reconstruction and notes on their ultrastructure. Zoological
Journal of the Linnean Society 116:71 – 84.
Wolburg-Buchholz, K., Greven, H. 1979. On the fine structure of
the spermatozoon of Isohypsibius granulifer Thulin 1928 (Eutardigrada) with reference to its differentiation, in: We˛glarska,
B., Ed., Second international symposium on tardigrades,
Kraków, Poland, 28 – 30, July 1977. Zeszyty Naukowe Uniwersytetu Jagiellonskiego, Prace Zoologiczne 25:191 – 197.
Wright, J. C. 1989a. The tardigrade cuticle. 2. Evidence for a dehydration-dependent permeability barrier in the intracuticle. Tissue and Cell 21:263 – 279.
Wright, J. C. 1989b. Desiccation tolerance and water-retentive mechanisms in tardigrades. Journal of Experimental Biology 142:
267 – 292.
Wright, J. C., Westh, P., Ramløv, H. 1992. Cryptobiosis in
Tardigrada. Biological Reviews of the Cambridge Philosophical
Society 67:1 – 29.
16
WATER MITES (HYDRACHNIDA)
AND OTHER ARACHNIDS
Ian M. Smith
David R. Cook
Biodiversity Section
ECORC, Central Experimental Farm
Agriculture and Agri-Food Canada
Ottawa, Ontario, Canada K1A 0C6
7836 North Invergordon Place
Paradise Valley, Arizona 85253
Bruce P. Smith
Biology Department, Ithaca College
Ithaca, New York 14850
I. Introduction to Arachnids
II. Water Mites (Hydrachnida)
A. Introduction
B. External Morphology and
Internal Anatomy
C. Life History
D. Habitats and Communities
E. Ecology
F. Collecting, Rearing and Preparation
for Study
I. INTRODUCTION TO ARACHNIDS
Members of the class Arachnida are the most
familiar representatives of the arthropodan subphylum
Chelicerata, an ancient lineage characterized by distinctive mouthparts with two pairs of appendages, the
anterior chelicerae and posterior pedipalps, and four
pairs of walking legs. Chelicerates first appear definitively in the fossil record as giant water scorpions
(Merostomata, subclass Eurypterida) in freshwater deposits from the Silurian. By the Devonian, eurypterids
had apparently disappeared but other chelicerate lineages were proliferating, such as horseshoe crabs
(Merostomata, subclass Xiphosura) in marine environments, and early derivative arachnid groups, including
mites, on land. Arachnids are the only group of chelicerates exhibiting extensive modern diversity. The
higher classification is still being debated, but arachnids
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
G. Taxonomic Keys to Genera of Water
Mites in North America
III. Other Arachnids in Freshwater Habitats
A. Other Mites (Acari)
B. Spiders
Acknowledgments
Literature Cited
are usually considered to be a taxonomic class comprising 11 extant subclasses, mostly terrestrial groups of
highly specialized predators of other arthropods. Only
two subclasses of modern arachnids are represented in
freshwater habitats, mites (Acari) which have diversified extensively to become arguably the most ubiquitous and adaptable clade of arthropods on Earth, and
spiders (Araneae) which have maintained a conservative way of life as essentially terrestrial predators.
Mites belonging to several unrelated groups are
commonly found in freshwater habitats. The true water mites comprise a taxon of actinedid acariform mites
which we refer to here as Hydrachnida (Hydrachnellae, Hydracarina, or Hydrachnidia of other authors).
Hydrachnida have flourished to become by far the
most numerous, diverse, and ecologically important
group of freshwater arachnids. Various families in certain suborders of Acariformes (namely, Actinedida,
551
552
Ian M. Smith, David R. Cook, Bruce P. Smith
Oribatida and Acaridida) and Parasitiformes (namely,
Gamasida) have independently invaded freshwater and
become adapted for living there. Compared to Hydrachnida, these groups are less diverse taxonomically,
usually less abundant, and relatively conservative in
morphology and habits. We summarize diagnostic and
ecological information for these mites and for the remarkable spiders of the genus Dolomedes at the end of
this chapter.
II. WATER MITES (HYDRACHNIDA)
A. Introduction
Water mites are among the most abundant and diverse benthic arthropods in many habitats. One square
meter area of substrate from littoral weed beds in eutrophic lakes may contain as many as 2000 deutonymphs and adults representing up to 75 species in
25 or more genera. Comparable samples from an
equivalent area of substrate in rocky riffles of streams
often yield over 5000 individuals of more than 50
species in over 30 genera (including both benthic and
hyporheic forms). Water mites have co-evolved with
some of the dominant insect groups in freshwater
ecosystems, especially nematocerous Diptera, and typically interact intimately with these insects at all stages
of their life histories.
We have made numerous additions and refinements to the information included in the first edition
of this chapter (Smith and Cook, 1991), and some extensive revisions. We have tried to avoid unnecessary
use of specialized acarological terms, and provide a
glossary of essential terminology needed for describing
morphological and anatomical features. Those wishing
to consult a more comprehensive account of mite
structure and function are referred to the appropriate
sections of the books by Cook (1974) and Krantz
(1978).
1. General Relationships
Hydrachnida, along with the enigmatic interstitial
Stygothrombidioidea and terrestrial Calyptostomatoidea, Trombidioidea, and Erythraeoidea, belong to a
remarkably diverse natural group of actinedid acariform mites, the Parasitengona. The complex and essentially holometabolous type of development of this
group is unique among Acari. Typically, after emerging
from the egg membrane, the hexapod larva seeks out
an appropriate host, and becomes an ectoparasite
which is passively transported while feeding on host
fluids. When fully engorged the larva transforms to the
quiescent protonymph (or nymphochrysalis). Radical
structural reorganization occurs during this stage,
giving rise to the active deutonymph which resembles
the adult in being octopod and typically predaceous,
but is sexually immature and exhibits incomplete sclerotization and chaetotaxy. The deutonymph feeds and
grows in size before entering another quiescent stage,
the tritonymph (or imagochrysalis). After completion
of metamorphosis during this stage, the mature adult
emerges (see Fig. 1).
Larval water mites can be distinguished morphologically from those of other parasitengones by having
two setae, rather than one, on the genu of the pedipalp.
Deutonymphal and adult water mites differ from all
other Acari in having glandularia on the idiosoma.
2. Origin
Water mites evolved from terrestrial stock, and hypotheses on their origin usually presume an ancestral
terrestrial parasitengone (Mitchell, 1957a; Davids and
Belier, 1979). A recently proposed alternative hypothesis (Wiggins et al., 1980; Smith and Oliver, 1986) suggests that the primitive parasitengone stock may have
been water mites resembling certain extant Hydryphantoidea. According to this postulate, water mites diverged from terrestrial ancestors with direct development, perhaps resembling extant Anystoidea, while
evolving the basic parasitengone life-history pattern as
a set of adaptations for exploiting spatially and temporally intermittent aquatic habitats.
The fossil record provides little insight into the
evolutionary history of water mites. The only reported
fossils that undoubtedly are water mites (Cook, 1957;
Poinar, 1985) are larval specimens from Tertiary deposits representing highly derived taxa. However, distributional data and host associations indicate that the
group originated no later than the Triassic or Jurassic
period.
Available morphological and behavioral data suggest that extant water mites are monophyletic
(Mitchell, 1957a; Barr, 1972; Cook, 1974; Smith and
Oliver, 1976, 1986), and that the major phyletic lineages of water mites, and possibly all Parasitengona,
were derived from hydryphantoidlike ancestors
(Mitchell, 1957a; Smith and Oliver, 1986).
3. Diversity and Classification
Well over 5000 species of water mites are currently
recognized worldwide, representing more than 300
genera and subgenera in over 100 families and subfamilies (Viets, 1987). Acarologists usually consider water
mites to be a taxon of intermediate rank between superfamily and suborder. Interestingly, the group appears to rival several orders of aquatic insects in diversity, and is comparable to some of them in age. Most of
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 1
553
Diagrammatic illustration of a generalized water mite history.
the genera and families are reasonably stable and
appear to approximate closely holophyletic lineages.
The families are conservatively grouped into seven
superfamilies. Three of these — Hydrovolzioidea, Hydrachnoidea, and Eylaoidea — probably represent natural groupings. The others — Hydryphantoidea, Lebertioidea, Hygrobatoidea and Arrenuroidea — are all
either paraphyletic or polyphyletic assemblages subject
to future revision. A comprehensive, detailed cladistic
analysis of extant fauna is required to permit construction of more stable and informative superfamily groupings. The phylogenetic studies that prove most useful in
understanding water mite evolution and developing a
more natural classification provide syntheses of information based on the morphology of all instars, behav-
ioral attributes, and life-history data (Mitchell, 1957a;
Cook, 1974; Smith, 1976a; Smith and Oliver, 1976,
1986).
The over 1500 species currently estimated to occur
in North America, north of Mexico, represent 136 genera in 69 subfamilies, 42 families, and all seven superfamilies. The fauna remains incompletely known at the
generic level, and revision of the family level classification is ongoing (see Smith, 1990b; Cook et al., 1998).
Species of over 25 new or unreported genera have been
discovered in North America since the publication of
Cook’s (1974) global review, an average of about one
each year (see Smith, 1976b, 1979, 1983b, c, d, g,
1989a – c, 1990a, 1991b, d, 1992b, 1998; Smith and
Cook, 1994, 1996, 1997, 1998a, b, 1999a, b; Gerecke
554
Ian M. Smith, David R. Cook, Bruce P. Smith
et al., 1999). Almost half of the species expected to
occur in North America are not yet named, and many
described species are known from only a few preserved
specimens in collections. The most diverse genera, such
as Sperchon, Torrenticola and Aturus in lotic habitats
and Piona and Arrenurus in lentic habitats, contain
100 or more nearctic species each (probably more than
250 species in the case of Arrenurus). Additional taxa
await discovery throughout the continent, especially in
springs and interstitial habitats. The least explored areas, such as southern Appalachia and the boreal and
arctic regions of northern Canada and Alaska, undoubtedly will yield some unexpected taxa when thoroughly studied.
inhabiting North America belong to these genera, and
descended from ancestors that either were present on
this continent when it separated from Eurasia during
the early Tertiary or immigrated from Eurasia with
their hosts via land bridges later in that Period.
4. Zoogeography
iv. Genera of North American origin Twenty-four
genera are currently known only from North America.
Some are probably endemics of recent origin while others have Tertiary relict distributions, suggesting that
they originated in Laurasia and may have undiscovered
members inhabiting poorly known areas of the Palearctic or Oriental Regions.
Distributions of water mite genera within North
America were dramatically influenced by climatic cooling and glaciation during the late Pliocene and Pleistocene. Genera which show evidence of recent adaptive radiation have re-established pan-continental
distributions, while those with relatively few species
have remained within more limited areas. Many
species groups came to be restricted to either temperate or boreal areas of the continent, as reflected to
some extent in the distributions of genera (see Table I).
As a result, typical “Tertiary-relict” distributions in, or
adjacent to, unglaciated refugia such as coastal California and Oregon, the Ozark Plateau, and parts of
the Appalachian and Rocky Mountains are exhibited
by some genera (Smith, 1989d, e, 1991c, 1992a, b) .
Other genera (e.g., Teutonia, Midea, Acalyptonotus)
have characteristic “boreal” or “arctic-alpine” distributions at higher latitudes and elevations. Distributional limitations are frequently correlated with habitat specializations. For example, many interstitial
genera have Tertiary-relict distributions because their
habitats were destroyed in most northern regions of
the continent by Pleistocene glaciation and will require
a long period of climatic stability to become re-established there. The sister species of many nearctic Tertiary relicts now inhabit similar areas in temperate
Asia, resulting in strikingly discontinuous distributions
for certain genera. In contrast, species adapted to cold
stenothermic pools inhabit subarctic areas, high mountains near permanent ice-fields, and temperate lowlands where groundwater springs create similar conditions. The sister-species of many stenophilic nearctic
a. Global Patterns Water mites occur throughout
the world, except Antarctica (Cook, 1974). All superfamilies, except Hydrovolzioidea, as well as many
early derivative families and subfamilies, are richly
represented in all zoogeographic regions. Knowledge
of global distribution patterns of water mite taxa has
been substantially improved by recent studies of
the fauna of austral regions, especially those by K.O.
Viets and D.R. Cook. The basic patterns can be explained as the result of vicariance due to plate tectonics (Cook, 1986, 1988; Smith and Cook, 1999c). Dispersal (along with hosts) between adjacent land
masses (e.g., Southeast Asia and Australia) or across
continents (e.g., Africa) significantly altered these patterns as more recent groups progressively displaced
ancient ones.
b. Nearctic Patterns Four major groups of genera
can be recognized among extant North American water
mites based on their apparent origins, reflecting successive historical stages in the development of the modern
fauna (see Table I).
i. Genera of Pangean origin Twenty-three earlyderivative, cosmopolitan genera exhibiting substantial
endemism in both the northern and southern hemispheres apparently originated in Pangea, and were represented in both Laurasia and Gondwanaland at the
time of their separation during the Jurassic.
ii. Genera of Laurasian origin Seventy-two genera with predominantly northern hemispheric distributions, exhibiting comparable diversity and endemism in
the Nearctic and Palearctic, probably originated in
Laurasia.
The genera of Pangean and Laurasian origin probably were distributed throughout Laurasia during the
late Cretaceous. The vast majority of species currently
iii. Genera of Gondwanan origin Seventeen genera with predominantly southern hemispheric distributions, exhibiting substantial diversity and endemism in
the Neotropics (Cook, 1980), are probably of Gondwanan or South American origin. Members of the
Gondwanan taxa invaded southern North America
from Central and South America after the Panamanian
isthmus appeared during the Pliocene.
16. Water Mites (Hydrachnida) and Other Arachnids
water mites live in northern Eurasia, yielding essentially circumpolar or boreal holarctic distribution patterns for these genera.
B. External Morphology and Internal Anatomy
Water mites exhibit the characteristic acarine body
plan comprising the gnathosoma, or mouth region, and
the idiosoma, or body proper, representing the fused
cephalothorax and abdomen. The standard morphological terminology proposed by Greandjean (see Robaux,
1974) has been widely adopted by most authors dealing
with prostigmatic mites, but until recently little effort
was made to apply this terminology to water mites. The
extreme plasticity of the adult exoskeleton during water
mite evolution has obscured many homologies with other
Prostigmata, and water mite workers have developed a
highly useful set of peculiar morphological terms for external structures of this instar. Early taxonomic studies of
larval water mites generated an additional set of special
morphological terms for the exoskeleton of this strongly
heteromorphic instar, and we used a modified version of
this terminology in the 1st edition. However, it has become evident that the relatively plesiotypical body plan
of larvae permits much of the standard idiosomal terminology of Grandjean to be used with confidence. We employ it here and indicate where appropriate the equivalence of the various terms in common use.
1. Morphology of Larvae
The short gnathosoma (Fig. 2) bears the stocky
pedipalps which have five free segments (trochanter, femur, genu, tibia, tarsus) that flex ventrally. The tarsus
of the pedipalp is relatively long and cylindrical in
some early derivative genera (eg: Wandesia, Thyas)
(Figs. 18, 22), but is typically reduced to a dome- or
button-shaped pad in most derivative groups (Figs. 76,
182). A highly modified thick, curved seta is present
dorsally at the end of the tibia (Fig. 2), the homolog of
the tibial “claw” which characterizes the pedipalp of
most terrestrial Anystoidea, Parasitengona and related
groups. The paired chelicerae (Fig. 3), each consisting
of a cylindrical basal segment and a movable terminal
claw, lie between the pedipalps.
The idiosoma is plesiotypically mainly unsclerotized, as in extant Hydryphantoidea. The dorsal integument (Fig. 27) bears a medial eye-spot, two pairs of
lenslike lateral eyes, four pairs of lyrifissures and the
following complement of setae: four pairs on the
propodosoma, two pairs of verticils, the internals (vi)
or anteromedials (AM) and externals (ve) or anterolaterals (AL), and two pairs of scapulars, the internals (si)
or sensillae (SS), and externals (se) or posterolaterals
(PL); eight pairs on the hysterosoma, three pairs in the
555
c-row (c1, c2 and c3), two pairs in the d-row (d1 and
d2), two pairs in the e-row, (e1 and e2) and one pair
from the f-row (f1). Ventrally (Fig. 28) the integument
bears the paired coxal plates, paired urstigmata (Ur)
laterally between plates I and II, the excretory pore
(EP), one pair of lyrifissures and the following complement of setae: three-to-six pairs on the coxal plates,
two in the 1-row (1a and 1b), zero, one or two in the
2-row (2b usually present, 2a and 2c usually absent)
and one or two in the 3-row (3a always present, 3b
usually absent), six pairs on the hysterosoma, one pair
of preanals (pa), two pairs associated with the excretory pore (ps1 and ps2), one pair from the f-row (f1)
and two pairs in the h-row (h1 and h2). The size,
shape, and position of these dorsal and ventral structures, and the degree of fusion of the sclerites associated with them, provide useful taxonomic characters.
The legs (Fig. 21) are inserted laterally on the coxal
plates, and in the plesiotypical condition have six movable segments, namely trochanter (Tr), basifemur
(BFe), telofemur (TFe), genu (Ge), tibia (Ti), and tarsus
(Ta), that articulate to permit ventral flexion. The segments have characteristic complements of setae and
solenidia (as in Figs. 6 – 8). The tarsi bear paired claws
and clawlike empodia terminally.
The soft-bodied condition, retained in all known larval Hydryphantoidea, apparently provides adequate support for the muscles used in locomotion on the surface
film. In all other major lineages, the dorsum (Figs. 3, 37,
56, 81, 144, 171) bears an extensive plate which incorporates the bases of the propodosomal setae and, in
some cases, one or more hysterosomal pairs. Similarly,
on the venter (Figs. 4, 38, 58, 83, 145, 172) the coxal
plates are enlarged and variously fused, and the excretory
pore plate (Figs. 5, 75, 109, 148, 176) is expanded to incorporate the bases of both pairs of excretory pore plate
setae. The coxal plates and excretory pore plate may also
bear various combinations of ventral setae.
Dorsal plates, and expanded coxal and excretory
pore plates, developed independently in Hydrovolzioidea and in most Eylaoidea to provide support
for muscles used in running and crawling on the surface film, and again in both Hydrachnoidea and the ancestral stock of the three “higher” superfamilies to anchor muscles used in swimming or crawling under
water. In Hygrobatoidea, fused coxal plates II III often have conspicuous lateral coxal apodemes (LCA),
medial coxal apodemes (MCA), and transverse muscle
attachment scars (TMAS) (Fig. 133).
Larvae of Hydryphantoidea and Eylaoidea retain
six movable leg segments (Figs. 21, 40), but those of all
other groups have the basifemoral and telofemoral segments fused (Figs. 6 – 8, 60). Larval Hydryphantoidea
also retain the most plesiotypical complement of setae
556
Ian M. Smith, David R. Cook, Bruce P. Smith
on the segments of the legs. Larvae of each of the more
derivative groups exhibit characteristic leg chaetotaxies
that reflect reductions from the plesiotypical complement. Certain ventral setae on the genua, tibiae and
tarsi are elongate and plumose in larvae adapted for
swimming (Fig. 149).
2. Morphology of Deutonymphs and Adults
The gnathosoma consists of the capitulum (or
gnathosomal base) and associated appendages, namely
the chelicerae and pedipalps (Fig. 200). The capitulum
is plesiotypically a simple, short channel, derived from
extensions of the pedipalpal coxae, leading to the oesophagus (Mitchell, 1962). A protrusible tube of integument connecting the capitulum to the idiosoma has
developed independently in several distantly related
genera (e.g., Rhyncholimnochares, Clathrosperchon,
Geayia). The paired pedipalps (Fig. 200, 227), inserted
on the capitulum, have both tactile and raptorial functions. In the plesiomorphic condition, the pedipalps
have five movable segments, namely trochanter, femur,
genu, tibia and tarsus, that are essentially cylindrical
and articulate to allow ventral flexion. The tibia bears
a thick, bladelike, dorsal seta distally in many ancient
genera (e.g., Hydryphantes, Tartarothyas, Pseudohydryphantes). As in larvae, this seta is the homolog of
the tibial “claw” of terrestrial relatives, and it often
makes the pedipalps appear chelate. In derivative
groups, other setae along with various denticles and tubercles may be elaborated to enhance the raptorial
function of the pedipalps. Segmentation of the
pedipalps is reduced by fusion in a few genera. A modification that has developed independently in various
groups of Arrenuroidea is the so-called uncate condition. In these genera, the tibia is expanded and produced ventrally to oppose the tarsus, permitting the
mites to grasp and hold slender appendages of prey organisms securely. The pedipalps are highly modified in
some interstitial genera (e.g., Fig. 426), presumably to
facilitate prey capture in confined hyporheic spaces.
The paired chelicerae lie in longitudinal grooves between the pedipalps on the dorsal surface of the capitulum. Plesiotypically they consist of a cylindrical basal
segment bearing a movable terminal claw. This cheliceral structure is designed for tearing the integument
of prey organisms, and is retained in nearly all derivative groups. Hydrachnidae are unique in having unsegmented and stilletoform chelicerae, an obvious adaptation for piercing the insect eggs upon which they feed.
The chelicerae are separate in all groups except Limnocharidae and Eylaidae where they are fused medially.
The most comprehensive comparative studies of water
mite mouthparts were published by Motas (1928) and
Mitchell (1962).
The idiosoma, or body proper, is plesiotypically
round or ovoid in outline, slightly flattened dorsoventrally, and mostly unsclerotized, as in certain extant
members of ancient genera such as Hydryphantes, Tartarothyas and Pseudohydryphantes. The dorsal integument (Figs. 224, 256) bears an unpaired medial eye,
paired lateral eyes that are usually enclosed in capsules,
paired preocular and postocular setae, and longitudinal
series of paired glandularia (six dorsoglandularia, five
lateroglandularia), muscle attachment sites (five dorsocentralia, four dorsolateralia), and lyrifissures (five).
Ventrally (Fig. 223) the integument bears the paired
coxal plates (fused into anterior and posterior groups
on each side), the genital field (comprising the gonopore, three pairs of acetabula, and paired genital
valves), five pairs of ventroglandularia (including coxoglandularia I between the anterior and posterior coxal
groups and coxoglandularia II behind the posterior
groups) and the excretory pore. As in larvae, these idiosomal structures provide a wealth of useful taxonomic
characters.
The legs (Figs. 200, 325) are inserted laterally on
the coxae, and plesiotypically articulate on a vertical
major axis and have six movable segments that articulate to permit ventral flexion. The segments are essentially cylindrical and have variable complements of setae. Though chaetotaxy of the legs provides a variety of
taxonomic characters in deutonymphs and adults, the
expression and position of individual setae are highly
variable within taxa. Consequently, the rigorous analysis of chaetotactic patterns that proves so useful in the
case of larvae is not practicable for later instars. The
leg tarsi plesiotypically bear paired claws terminally.
The plesiotypical soft-bodied condition is retained
in early derivative genera adapted for walking on the
substrate (e.g., Thyas) or swimming in shallow habitats
such as seepage pools or temporary pools (e.g.,
Hydryphantes, Pseudohydryphantes). Walking forms
have relatively short, stocky leg segments and setae,
whereas swimmers have longer segments bearing fringes
of slender swimming setae. As pointed out by Mitchell
(1957a, b, 1958, 1964b), the soft integument of these
mites permits the highly precise local control of body
shape and internal pressure that is needed to produce
the walking and swimming movements of the legs.
3. Evolution of Adult Exoskeleton
The major lineages of water mites apparently differentiated from ancestral stock that resembled extant
soft-bodied Hydryphantoidea (Mitchell, 1957a, b;
Smith and Oliver, 1986), primarily through development of apotypical larval behavioral patterns and life
histories. Hydrachnidae and Eylaidae became specialized for exploiting standing water habitats, especially
16. Water Mites (Hydrachnida) and Other Arachnids
temporary pools, whereas Hydryphantoidea, Lebertioidea, Hygrobatoidea, and Arrenuroidea ultimately
diversified and radiated in a wide range of habitats. In
each of these four groups, and presumably in Hydrovolzioidea, sclerotization of the integument came about
as soft-bodied ancestral stock began to invade new
habitats and diverge.
Adaptation to habitats such as seepage areas and
springs or substrates in streams required a change in locomotor habits from walking or swimming. Mites
adapting to moss mats and wet litter habitats developed hydrophilic integument which draws a film of
water over the dorsal surface, creating sufficient downward force to press the body to the substrate and prevent efficient walking (Mitchell, 1960). Groups invading streams evolved a wedge-shaped body designed for
negotiating confined spaces to avoid exposure to water
turbulence and strong currents. Locomotion in both of
these types of habitats necessitated evolution of a
crawling gait. This change involved a shift in orientation of the major axis of the legs, shortening and thickening of leg segments and setae, enlargement of tarsal
claws, and development of stronger, more massive
muscles to control leg movements. Expansion of coxal
plates and sclerites associated with glandularia, setal
bases, and the genital field, along with sclerotization of
the dorso — and laterocentralia, occurred to provide
rigid exoskeletal support for these muscles. Fusion of
these sclerotized areas led to development of complete
dorsal and ventral shields in adults of certain groups.
Multiple invasions of helocrene, rheocrene, and lotic
habitats gave rise to a diverse array of dorsoventrally
flattened, sclerotized, crawling groups of Hydryphantoidea, Lebertioidea, Hygrobatoidea and Arrenuroidea.
During the early evolution of each of these lineages
certain groups became adapted for exploiting interstitial habitats in subterranean waters and the hyporheic
zone of rheocrenes and streams. These mites tended to
lose eyes and integumental pigmentation, and required
further stream-lining of the body to facilitate locomotion in interstitial spaces. Consequently, soft-bodied
forms adopted a vermiform shape, while sclerotized
mites became extremely compressed laterally or
dorsoventrally. The crawling mode of locomotion became secondarily modified in several groups of wellsclerotized interstitial genera (e.g., Neomamersa, Frontipodopsis, Chappuisides) to permit rapid and agile
running in hyporheic and ground water habitats.
By late Pangean times, evolution within water
mites apparently had produced basic communities of
essentially soft-bodied species living in temporary and
permanent standing water, and partly to fully sclerotized species living in emergent groundwater, lotic, and
interstitial habitats. Subsequent phylogeny of
557
Hydryphantoidea, Lebertioidea, Hygrobatoidea, and
Arrenuroidea during late Mesozoic, Tertiary, and Quaternary times appears to have followed a recurring pattern of extended periods of gradual evolution leading
to specialization within particular habitats, punctuated
by episodes of relatively rapid and dramatic diversification. Divergence of many of the clades that originated
during this time appears to have been correlated with
successful invasion of new habitats. Repeated habitat
diversification within the major clades of these four superfamilies, in response to opportunities provided by
the explosive evolution of their primary host groups
and by prolonged geological and climatic instability, resulted in several parallel and convergent trends in body
sclerotization. Members of different clades developed
superficially similar sclerite arrangements in adapting
to lotic or interstitial habitats (e.g., species of Diamphidaxona and certain Axonopsinae); others underwent
homoplastic reduction or loss of sclerites in secondarily
invading lentic habitats (e.g., species of Forelia and Piona). Finally, some groups retained extensive sclerotization while becoming adapted for swimming in standing water (e.g., certain species of Axonopsis and
Mideopsis). Consequently, modern communities are
highly heterogeneous phylogenetically and morphologically, with each monophyletic component exhibiting
unique exoskeletal adaptations for living in the particular habitat.
Evolutionary trends involving other readily observed and taxonomically useful characters — such as
position and number of genital acetabula, positions of
glandularia and lyrifissures, number and arrangement
of setae on the body plates, and modification of claws
on the tarsi of the legs — are still not adequately explained. For example, although the genital acetabula
are plesiotypically borne in the gonopore in most
Hydryphantoidea and Lebertioidea, they are either
fused with the genital flaps or incorporated into acetabular plates flanking the gonopore in most taxa of other
superfamilies. In addition, the number of acetabula has
proliferated from the plesiotypical complement of three
pairs several times independently in each of the six
largest superfamilies. Alberti (1977, 1979) has recently
proposed that acetabula in water mites may function as
an osmoregulatory chloride epithelium, and Barr
(1982) concluded that analysis of morphological data
for early derivative taxa supports this hypothesis.
However, until the function of acetabula is clearly understood on the basis of well-designed experiments, the
significance of the trend to increasing number of acetabula will remain speculative.
The often striking and elaborate color patterns of
adult water mites present an intriguing enigma. Members of most ancient clades (e.g., Hydrachnoidea, Ey-
558
Ian M. Smith, David R. Cook, Bruce P. Smith
laoidea, Hydryphantoidea) have colorless integument
but are red, like terrestrial parasitengones, due to the
presence of pigment granules which appear to be distributed throughout the body. Members of other superfamilies, except for taxa in interstitial habitats, exhibit
highly distinctive patterns resulting from symmetrically
arranged concentrations of pigment granules of various
colors. In soft bodied taxa the pigments are located beneath the integument, but in sclerotized groups they are
incorporated into the plates. Members of well-sclerotized taxa of Lebertioidea, Hygrobatoidea, and Arrenuroidea may be various shades of red, orange, yellow, green, or blue, and the dorsal shield frequently
exhibits an intricate pattern combining several contrasting colors. The dorsal patterns of these mites can
readily be interpreted as disruptive camouflage, but the
adaptive value of their bright colors is more difficult to
understand. The evolution of a wide range of highly
colored pigments in these taxa certainly suggests that
distinctive coloration confers some selective advantage.
Though protection from predation may be part of the
explanation, it is tempting to speculate that these mites,
despite their apparently simple eye structure (see below), may detect and use color as one of several cues
for recognizing conspecific individuals. Research is
needed to determine the extent of their ability to detect
and respond to the often subtle differences in color that
distinguish closely related taxa.
4. Internal Anatomy
The organ systems of water mites occupy the hemocoel, bathed by hemolymph which is circulated by
movements of the body musculature (Schmid, 1935;
Bader, 1938; Mitchell, 1964b). As is typical for arthropods, there is no closed circulatory system.
a. Digestive System Digestion of food materials
begins pre-orally, and only fluids are ingested. Food is
drawn into the mouth (or buccal cavity) by the muscular pharynx, then passed through the tubular esophagus to the lobed midgut (Bader, 1938) where digestion
and absorption occur. Undigested material accumulates
as insoluble particles in a posterodorsal lobe of the
midgut. All lobes of the midgut end blindly, and there
is no connection between the gut and the excretory
pore.
b. Excretory System This system consists of a
large, thin-walled excretory tubule, apparently derived
from the primitive hindgut, that lies dorsal to, and in
close contact with, the midgut. Waste products are absorbed from hemolymph, and stored in the excretory
tubule as insoluble, whitish or yellowish crystals of unknown chemical composition. When filled with crys-
talline material the excretory tubule may be visible
through the dorsal integument as a “T”- or “Y”shaped structure. The excretory tubule connects ventrally with the excretory pore, and is evacuated periodically by pressure generated through movements of the
body muscles.
Osmoregulation is apparently accomplished by the
urstigmata in larvae, and by the genital acetabula in
deutonymphs and adults (Barr, 1982). Both of these
structures have porous caps which apparently provide
an ample surface area of chloride epithelium for maintaining water balance.
c. Respiratory System Many groups of water
mites retain paired stigmata located between the bases
of the chelicerae in all active instars. Plesiotypically, the
stigmata lead to tracheal trunks which anastomose repeatedly into tracheolar tubules extending to all parts
of the body. However, they appear to be nonfunctional
in deutonymphs and adults, and respiration occurs by
diffusion through the integument. In adults of many
large species inhabiting standing water, a network of
closed tubes of tracheolar dimensions lying beneath the
integument transports gases to and from the tracheae
that lead to internal organs (Mitchell, 1972). Heavily
sclerotized mites have the body plates well supplied
with regularly arranged “pores” that permit diffusion
of gases between the tracheolar loops and surrounding
water through areas of thin integument. In some
species, there are regions of tracheal anastomosis lateral to the brain (Wiles, 1984).
d. Neural System The so-called brain, a fused, undifferentiated central ganglionic mass, surrounds the
oesophagus. Nerve trunks dorsal to the oesophagus
lead to the anterior sense organs and mouthparts, while
ventral trunks lead to the legs and genital region.
The lateral eyes appear to be the primary lightsensing structures in water mites. There are typically
two pairs, with the eyes of each side located close together and often enclosed in lenslike capsules that may
lie above or beneath the integument. In postlarval instars of Eylaoidea, the eyes are borne on a medial sclerite. Many Hydryphantoidea also have a medial eye
spot between the lateral eyes, but this structure is absent in most members of the other superfamilies. The
eyes seem to function as ocelli, permitting the mites to
detect the intensity and direction, and, at least in some
taxa, the wavelength of incident light. Experiments
with species of Unionicola have demonstrated that
some species respond differentially to various wavelengths (Dimock and Davids, 1985) and that the response of the mites can be influenced by chemicals produced and released by their mollusc hosts (Roberts
16. Water Mites (Hydrachnida) and Other Arachnids
et al., 1978). Retinal development, though rudimentary, has been reported for a variety of taxa (Lang,
1905).
Members of the various taxa have highly characteristic complements of setae on the idiosoma and appendages. Setae are the main tactile receptors, and
many of them have also assumed secondary functions
in locomotion, feeding, or mating. All water mites have
specialized tactile organs, called glandularia, distributed in pairs on both the dorsum and venter of the
body. Each glandularium consists of a small, gobletshaped sac, or “gland”, with a tiny external opening,
and an associated seta. The “gland” contains a milky,
viscous fluid which is ejected abruptly when the seta in
stimulated. This substance quickly stiffens to a sticky
gel after it comes in contact with water, and apparently
evolved as a deterrent to attackers. In males of Arrenurus, however, material secreted by the glandularia
seems to function also in cementing the body of the female in place during copulation.
The five pairs of lyrifissures, which are arranged in
series on the dorsum and venter of the body are
thought to be proprioceptors (Krantz, 1978). In addition, the distal segments of the pedipalps and legs are
well supplied with a variety of specialized setiform
structures, apparently derived from outgrowths of the
integument, which function as chemoreceptors. The
most conspicuous of these organs are the solenidia
which adorn the dorsal surfaces of the genua, tibiae,
and tarsi of the legs, and the tarsi of the pedipalps.
e. Reproductive System Males have paired testes
and vasa deferentia which lead to an elaborate ejaculatory complex consisting of a series of membranous
chambers attached to a sclerotized framework (Barr,
1972). Positioned immediately above the genital field,
the ejaculatory complex functions as a syringelike organ for compacting masses of spermatozoa, assembling
spermatophores, and expelling them from the genital
tract through the gonopore.
Females have paired ovaries that are more or less
fused, and paired oviducts that also fuse to form a single duct leading to the genital chamber within the
gonopore. Paired spermathecae, for storing spermatozoa picked up in spermatophores, flank the genital
chamber and are connected to it by short ducts.
C. Life History
The basic life-history pattern of water mites (Fig.
1) was first correctly inferred by Wesenburg-Lund
(1918) and has been confirmed by subsequent studies
on a wide range of taxa (see Smith and Oliver, 1986,
for a summary of relevant literature). The first well-
559
documented life-history studies were of palearctic
species, especially members of the genus Arrenurus, investigated by Münchberg (1935a, b). Subsequently,
Mitchell (1959, 1964a) dealt intensively with a number
of nearctic species of Arrenurus, and Böttger (1962,
1972a, b) described life cycles of representative
palearctic species of Hydrachna, Limnochares, Eylais,
Limnesia, Unionicola, Piona, and Arrenurus in considerable detail. A meticulously comprehensive account of
the life history of Hydrodroma despiciens (Müller)
(family Hydrodromidae) was published by Meyer
(1985).
Water mites in temperate latitudes typically live for
just over one year, most of which is spent in the deutonymphal and adult stages. Most species are univoltine, with long-lived female adults producing multiple
clutches of eggs. This tactic apparently minimizes the
risks associated with the parasitic larval stage. In temperate latitudes, hosts are usually seasonally limited
and parasitism occurs mainly in late spring and early
summer. Larvae of most taxa spend no more than several days on their hosts and engorgement results in
modest growth. Deutonymphal water mites typically
feed and grow throughout the summer and adults appear in late summer or early fall and mate almost immediately. Inseminated females remain in obligative reproductive diapause until the following spring. This
type of life history occurs in a wide range of water mite
clades associated with host groups that are
holometabolous and have aerial imagoes.
Several derivative life-history strategies occur
among water mites, representing variations on this basic pattern (Smith and Oliver, 1986; Smith, 1988,
1999). One of these involves elimination of larval feeding and host association altogether, thus avoiding risk
of not encountering a host but foregoing host-mediated
dispersal (Smith, 1998). This has happened at least 20
different times in unrelated genera representing a wide
diversity of families. Comparison of closely related
species pairs with, and without, larval parasitism
shows that loss of larval parasitism correlates with accelerated maturity and metamorphosis to adult at a
smaller body size (Smith, 1998). Removal of the seasonal limitations of host association in species with
nonparasitic larvae appears to favor a shift to a multivoltine life history. Several clades exhibit a third strategy by greatly extending the association with hosts, in
some cases for several months, through the larval and
protonymphal stages. Species in these groups typically
undergo remarkable growth during the larvae stage, increasing in volume by up to 700 times their original
size (Davids, 1973). Growth of these mites as deutonymphs is much more modest, and this stage often
lasts only a few days or weeks. Extended larval associa-
560
Ian M. Smith, David R. Cook, Bruce P. Smith
tion with hosts is exhibited by all species of Hydrachnoidea and Eylaioidea (excluding Limnochares), and
also by atypical members of certain other taxa, such as
Hydryphantes tenuabilis Marshall (Lanciani, 1971a).
All known examples of extended association involve
larval parasites of aquatic Hemiptera and Coleoptera,
presumably reflecting the longevity, large size and
mainly aquatic existence of these hosts. This type of life
history allows at least some species to adopt multivoltine life cycles (Davids, 1973). There are interesting
variations on these major life history patterns among
species within genera (Lanciani, 1971b; Cook et al.,
1989).
1. Egg
Eggs are typically laid in masses in a gelatinous
matrix and attached to plants, wood particles, or
stones, but females of some species, such as Midea expansa Marshall, may scatter them individually on the
substrate. Females of the genus Hydrachna use a
uniquely developed, elongate ovipositor to lay eggs individually in stems of aquatic plants, and those of certain species of Unionicola use short stylets to oviposit
in the tissues of sponges or mussels. The most comprehensive morphological study of water mite eggs was
published by Sokolow (1977). Detailed accounts of
oviposition behavior have been recorded by Crowell
(1960), Ellis-Adam and Davids (1970) and Davids
(1973).
Egg production varies greatly among species. For
example, females of Eylais discreta Koenike may produce individual clutches of 1000 – 2000 eggs, with one
individual able to produce over 13,000 eggs in a 3month period (Davids, 1973). On the other extreme,
species of Thyas, Hygrobates, Piona and Arrenurus
that forego larval feeding typically produce 2 – 5 eggs
per clutch (Smith, 1998). Most water mite species typically produce about a dozen-to-several hundred eggs in
a clutch.
There is an evident trade-off between clutch size
and egg size, probably related to risks to the larval instar and the proportion of total growth that occurs
during this stage. An analysis of this trade-off in a
group of 27 species of Arrenurus resulted in two evolutionary trajectories defined by egg size, clutch size, and
female size: one representing species parasitic on adult
nematocerous flies and the other species parasitizing
adult Odonata (Cook et al., 1989).
Large, communal clutches of eggs have been observed in several situations. Patches of eggs of Eylais
euryhalina Smith in western Canada frequently cover
areas of more than 100 cm2 on plant or rock substrates. Females of Arrenurus pseudosuperior Cook
produce communal egg masses entirely covering sub-
merged oak leaves in central Canadian lakes. The
stimulus to lay eggs communally may stem from a
shortage of oviposition sites in nature, although there
may be selective advantage if either eggs are distasteful
or their sheer numbers exceed what local predators can
consume.
Within the egg membrane, water mites pass
through a transitory prelarval instar (Meyer, 1985) and
then usually develop rapidly and directly into larvae.
Fully formed individuals can be observed moving
within the egg membrane just prior to emerging 1 – 3
weeks after oviposition. Arrested larval development
has been reported in certain species of Unionicola
(Mitchell, 1955), Teutonia (Smith, 1982), Utaxatax
(Smith, 1982) and Laversia whose larvae do not become active and emerge until at least six months after
eggs are laid.
Female mites presumably possess adaptations for
selecting appropriate oviposition sites.
2. Larva
The vast majority of water mite species are parasitic on imaginal insects as larvae (reviewed by Smith
and Oliver, 1976, 1986, and summarized in Table I),
with the host providing nutrition as well as dispersal.
Growth on the host is modest in many clades, but apparently essential for development as there are no
records of purely phoretic associations. Mitchell (1966,
1969a) theorized that the probability of success for larval water mites can be calculated as the product of the
probabilities of discovering a host, attaching at an appropriate site on the host, completing engorgement on
the host and detaching from the host in a suitable habitat. The risk of failure can be high. Collins (1975) estimated that more than 75% of larval Wandesia (Partnuniella) thermalis Viets fail to locate a host in a
system in which the distribution of hosts is extremely
clustered and unpredictable (Collins et al., 1976).
a. Evolution of Parasitic Associations
i. Host selection After emerging, larvae must
quickly locate hosts that both provide adequate sites
for attachment and feeding and regularly visit habitats
that are suitable for postlarval development. Insect
hosts provide water mites with both the source of nutrition necessary for larval growth, and their primary
dispersal mechanism. Host associations of larval water
mites were reviewed by Smith and Oliver (1976, 1986),
and are summarized in Table I.
Larvae of Hydrovolzia, and most genera of
Eylaoidea and Hydryphantoidea, rise to search for
potential hosts on the surface film, exhibiting plesiotypical terrestrial behavior. In contrast, larvae of
16. Water Mites (Hydrachnida) and Other Arachnids
Acherontacarus, Rhyncholimnochares, Wandesia, Hydrachna, Lebertioidea, Hygrobatoidea, and Arrenuroidea begin to swim in the water column or crawl
on the substrate as fully adapted aquatic organisms.
This apotypical behavior developed independently several times, within the superfamilies Hydrovolzioidea,
Eylaioidea and Hydryphantoidea, in ancestral Hydrachnidae, and again in ancestors of the other three
superfamilies. The development of fully aquatic larvae
can best be understood as an adaptation enhancing the
probability of success in locating hosts and, in the
higher water mites, facilitating preparasitic attendance
of hosts during their final preadult instar.
Surface dwelling larvae are essentially terrestrial,
and typically can locate hosts only when they visit or
pass through the surface film. Larvae of Limnochares
americana Lundblad are unusual in being able to
search for hosts up to 50 cm. away from the water surface. Opportunities of these mites to contact hosts are
transitory events occurring irregularly in space and
time, and are strongly influenced by specific aspects of
behavior within a host population. For example, parasitic larvae of Thyas barbigera Viets are found only on
parous female mosquitoes and those of Limnochares
americana are found predominantly on territorial male
dragonflies (Smith and Cook, 1991; Smith, 1999).
Some water mites with terrestrial larvae overcome the
risks of failure by utilizing a relatively broad range of
host organisms and producing large numbers of eggs.
For example, larvae of species of Hydrovolzia and certain genera of Hydryphantidae apparently exploit a
wide range of hosts, including insects that are only casual visitors to the habitat of the mites. This strategy
presumably results in considerable wastage, as larvae
attaching to hosts that do not return to suitable mite
habitats will die.
Larvae of most Eylaoidea (Eylais, Piersigia, Neolimnochares, some Limnochares) and various
Hydryphantidae have developed strong specificity for
particular groups of insect hosts that live on the surface
film (e.g., Hydrometridae, Gerridae), regularly visit it
to replenish air supplies (e.g., aquatic Hemiptera and
Coleoptera), or pass through it just before ecdysis and
regularly revisit it after emergence (e.g., Odonata, Trichoptera, Diptera). This strategy improves the probability that a larva finding a host will be returned to an
appropriate habitat, but limits the availability of potential hosts. These mites apparently compensate for the
high risk of failure in finding hosts by producing many
hundred eggs per female.
Species with terrestrial larvae are generally limited
to relatively shallow water microhabitats, given that
these larvae must swim or crawl to the surface in order
to search for potential hosts. Enclosure experiments by
561
one of us (B.P. Smith) indicate that the likelihood of
water striders of the genus Gerris being parasitized by
larval Limnochares aquatica Linnaeus is strongly dependent on water depth. Differences in rates of infestation are substantial, even between insects caged over
0.5 m of water and those caged over 1.5 m.
In contrast, aquatic larvae have access to potential
hosts throughout their aquatic existence and can actively seek them in the water column or substrate. Larvae of Hydrachnidae exploit virtually the same range
of hosts as Eylaidae, but locate them opportunistically
beneath the surface film with most species exhibiting
no evidence of preparasitic attendance. Members of
certain Anisitsiellidae and Teutoniidae (Lebertioidea),
as well as Krendowskiidae and Arrenuridae, apparently
also seek their hosts above the substrate, but have
evolved preparsitic attendance of hosts during their
penultimate instar prior to ecdysis. Their larvae parasitize tanypodine Chironomidae, Culicidae, Ceratopogonidae, Chaoboridae and Odonata, all groups that
have the final aquatic instar active in the water column
below the surface film.
Mites locating their hosts in the water column apparently exploit them more efficiently than do those
with terrestrial larvae, as their females tend to lay
smaller, though still substantial, numbers of eggs. For
example, females of Hydrachna conjecta (Koenike) lay
only about 10% as many eggs as those of Eylais discreta Koenike, yet these species co-occur in nature and
parasitize the same species of corixids (Davids, 1973).
Larvae of Rhyncholimnochares and Wandesia are
also aquatic and apparently independently evolved
adaptations to locate their hosts in the substrate. Larvae of Rhyncolimnochares are subelytral parasites of
adult elmids that remain in water. Those of Wandesia
attend nymphal stoneflies before transferring to the
emerging adults and becoming parasites. Interestingly,
larval stygothrombidiids exhibit similar behavior in exploiting the same hosts. Larvae of the remaining families of Lebertioidea, Hygrobatoidea, and Arrenuroidea
locate hosts as final instar larvae or pupae in cases or
burrows in the substrate. The hosts, various Chironomidae or Trichoptera, construct cases or retreats where
they remain during prepupal and pupal stages. These
mites have developed suites of larval adaptations to exploit this sedentary behavior, greatly increasing their
probability of successfully locating and associating
with appropriate hosts. Larvae in all of these clades attend the host during the preparasitic phase, clinging to
the integument of the pupa until it rises to the surface.
They typically then transfer to the imago as it emerges,
embed their chelicerae, and enter the parasitic phase.
Individual females in these groups lay relatively small
numbers of eggs (often less than 10), indicating that the
562
Ian M. Smith, David R. Cook, Bruce P. Smith
larvae efficiently find hosts with a high probability of
returning to a suitable habitat.
The development of fully aquatic larvae had important consequences each time it occurred during water mite evolution. In particular, it was a crucial preadaptation for ancestors of the various groups of
Lebertioidea, Hygrobatoidea, and Arrenuroidea that
became specialized in exploiting nematocerous Diptera
as hosts. The origin and evolution of modern genera
and families of these mites apparently followed the explosive radiation of the Nematocera, and especially
Chironomidae, during the Jurassic and early Cretaceous. Refinement of their elegant adaptations for parasitizing these flies permitted the so-called higher water
mites to co-evolve with them, and to exploit their potential for dispersal, invasion of new habitats, and
rapid evolution.
ii. Site selection Larvae of many subfamilies of
water mites exhibit strong selectivity for attaching to
particular sites on the body of the host (Smith and
Oliver, 1976, 1986; Oliver and Smith, 1980), as shown
in Table I. These preferences appear to illustrate another
adaptive process that has permitted mites to coevolve
successfully with nematocerous flies by minimizing interference with the dispersal potential of the host.
Larvae of Hydrovolzia and certain genera of
Hydryphantidae that parasitize Hemiptera, Plecoptera
and Trichoptera (e.g., Protzia, Wandesia), seem to attach rather indiscriminately to various parts of the
host’s body. Those of all other groups are more selective. Larval Eylais are essentially terrestrial organisms
and select attachment sites bathed by the host’s air
supplies, under the elytra of aquatic bugs and beetles
(Fig. 427). Significantly, the fully aquatic larvae of
Hydrachna are able to utilize a much wider range of
sites on the same hosts (Fig. 428).
Larval hydryphantoid mites that parasitize Diptera
tend to attach to thoracic sites, exhibiting apparently
plesiotypical behavior. Larvae of many early derivative
groups of Lebertioidea, and most Arrenuroidea parasitizing Diptera, also prefer thoracic sites. These larvae
tend to be relatively large, and only a few individuals
can share a single host. When more than one of these
larvae attach to the same part of the host’s thorax they
are usually symmetrically arranged on either side of the
body. Larvae of some species of Sperchon, Lebertia,
Oxus and Torrenticola can use the anterior abdominal
segments of hosts when thoracic sites are already
occupied.
In virtually all Hygrobatoidea (except Limnesiidae
which may not be closely related to other taxa in the superfamily) and a few families of Arrenuroidea (eg.,
Mideidae, Momoniidae) larvae attach exclusively to ab-
dominal sites on their hosts, usually nematocerous flies
but in some cases caddis flies. This apotypical behavior
correlates with a marked trend to smaller size, allowing
several larvae to share each attachment site on a host.
These small larvae occupying abdominal attachment sites
can exploit even the smallest species of Chironomidae as
hosts without preventing them from flying and dispersing
(Fig. 430). The relatively small larvae of various species
of Arrenurus (sensu stricto) show preferences for either
thoracic or abdominal sites on odonate hosts (Fig. 429).
The duration of the parasitic phase of larval development varies considerably. Larvae of certain species in
early derivative genera such as Limnochares and
Hydryphantes retain their mobility and can transfer
from one instar to the next of the same host individual.
Larval Limnochares aquatica engorge gradually by feeding on successive host instars, but larvae of
Hydryphantes tenuabilis exhibit true preparasitic attendance and delay feeding until the host reaches adulthood. This stategy of tracking hemimetabolous hosts
through several molts may reflect the plesiotypical condition in water mites. Larval Hydrachna and Eylais
remain attached to the same adult bug or beetle
throughout the parasitic phase, for several weeks or
more. Larvae of species in these families that are
adapted to temporary pools often remain on hosts
throughout the dry phase of the habitat, and only
engorge after a period of arrested development that may
last as long as 10 months. Larvae of species parasitizing
nematocerous Diptera or other aerial insects typically
engorge rapidly and mature within several days. Fully
fed larvae enter the quiescent protonymph stage.
The growth and dispersal functions of the larva
have both strongly influenced the evolution of host associated behavior in water mites, with different results
in the various major clades. Larvae of early derivative
genera such as Hydrachna, Eylais, and certain
Hydryphantidae grow substantially, increasing several
hundred-fold in size in some cases, while feeding on
their relatively long-lived hosts. Larvae of many relatively recently evolved groups associated with shortlived insects, especially nematocerous Diptera, still engorge on fluids from the host but increase more
modestly in size in the process. The primary strategic
role of the larval instar in the life history of most
Hydryphantoidea, Lebertioidea, Hygrobatoidea and
Arrenuroidea appears to be dispersal. Most feeding and
growth occur during the deutonymph and adult stages
in these mites.
The larva may be suppressed as an active instar in
a few unrelated species in widely divergent genera (e.g.,
Thyas, Lebertia, Limnesia, Hygrobates, Piona, Axonopsis and Arrenurus), apparently permitting the
mites to accelerate development and maximize ex-
16. Water Mites (Hydrachnida) and Other Arachnids
ploitation of optimal conditions (see Smith and Oliver,
1986; Smith, 1998). In these cases, larvae either remain
within egg membranes or emerge for a brief period of
activity before entering the protonymph stage. Species
known to forego a parasitic phase appear to be isolated
and ephemeral evolutionary experiments (Smith and
Oliver, 1986; Smith, 1998). Recent studies have detected relatively high levels of morphological divergence and lower levels of heterozygosity within populations of apparently recently diverged species lacking
parasitism when compared with closely related species
that retain parasitic larvae (Bohonak, 1998). We conclude that although elimination of parasitism confers
short-term advantage in larval survival and potential
for population growth, host-mediated dispersal is essential for long-term persistence of a lineage.
b. Ecological Aspects
i. Emergence of larvae In permanent water habitats at north temperate latitudes, most species of water
mites overwinter primarily as inseminated females and
there appears to be a necessary time interval in the
spring for oviposition and development of larvae to
proceed before parasitism can commence. Consequently, aquatic insect species whose adults emerge in
early spring are rarely parasitized by water mites, even
within groups that are otherwise suitable hosts. Interestingly, some of these early emerging species show
high susceptibility to parasitism under laboratory conditions. For example, although only a small proportion
of Aedes communis and Aedes punctor mosquitoes
carry larval Arrenurus angustilimbatus and Arrenurus
kenki in nature, they prove to be more susceptible to
parasitism in choice experiments than the species typically utilized as hosts by these mites in the field (Smith
and McIver, 1984a, b). A few mite species that apparently overwinter as eggs or larvae are adapted to exploit hosts that emerge in early spring. For example,
larvae of Huitfeldtia rectipes Thor are found regularly
on imagoes of early season species of Chironomus
emerging from oligotrophic lakes in eastern Canada
throughout the ice-free period beginning in mid-April.
Available evidence suggests only loose synchrony between life cycles of mites and their host species, even
when host specificity is strong (Smith and McIver,
1984a, c; Smith and Cook, 1991). In the case of Coquillettidia perturbans mosquitoes in Florida parasitized by larval Arrenurus danbyensis and Arrenurus
delawarensis, peak infestation did not closely correlate
with peak abundance of hosts and there was no close
seasonal correspondence between population densities
of the two mite species although both are dependent on
the same host (Lanciani and McLaughlin, 1989).
563
ii. Seeking and locating hosts After emerging,
larvae of species with a parasitic phase immediately begin host-seeking. The possible period of survival for
free-living larvae in the preparasitic phase ranges from
4 days to 6 weeks (Smith, 1988), but is about 1 week
for most species. Larval Arrenurus remain active and
exhibit preparasitic attendance over a period of 2
weeks in the laboratory, but have the greatest chance to
locate and infect a host during the first few days. Larvae older than 1 week rarely successfully attach to
hosts, although those of some cold-adapted species apparently can remain infective for longer periods.
Some authors have suggested that water mite larvae find hosts by accidental contact, but close observation of both terrestrial and aquatic larvae reveals noticeable behavioral changes when in proximity to hosts.
Terrestrial larvae seem to use visual, tactile and chemical cues to locate hosts at the surface film. Those of
several Hydryphantidae have the hind legs modified for
jumping and leap toward objects positioned above
them, an obvious adaptation for encountering hosts on
the surface film. Larval Limnochares aquatica exhibit
questing behavior when within several millimeters of
water striders, involving hesitation and reaching upward with the first pair of legs. Details of the sensory
mechanisms used by aquatic larvae to locate hosts remain unclear, but there is evidence that both chemical
and tactile cues are important (Böttger, 1972b). Larvae
of Hydrachna often extend the gnathosoma when approaching a host before actual contact is made. Larval
Arrenurus are attracted to mosquito pupae and slow
their swimming just prior to contact (Smith and
McIver, 1984b). These larvae swim in tight circles, as if
attempting to orient to some cue, before making a direct approach (Page and B.P. Smith, unpublished data).
iii. Recognizing host Larvae of water mite
species are typically able to exploit a range of host
species. Some species with aerial larvae have remarkably broad host spectra encompassing several orders of
insects, whereas species with preparasitic attendance
will normally be limited to species within a subfamily
or family of host insects. Commonly, one or two host
species carry the majority of parasitic larvae of a given
mite species in a particular habitat.
Although co-occurrence in space and time set the
absolute boundaries for host utilization, larval mites
also actively select among potential host species. In laboratory experiments, larval mites of diverse genera,
such as Hydrachna, Eylais, Limnochares and Arrenurus exhibit preferences when exposed simultaneously to
hosts of various species (Smith, 1977; Smith and
McIver, 1984b). Choice of host by mite larvae appears
to be influenced by differences in the intensity of cues,
564
Ian M. Smith, David R. Cook, Bruce P. Smith
and larval mites favor certain individuals within a host
population while rejecting others (Smith and McIver,
1984b). In some instances, hosts react strongly to the
presence of mite larvae with avoidance behavior or vigorous grooming activity (Forbes and Baker, 1990;
Smith and McIver, 1984b), and there may be differences in intensity of parasite avoidance among host
species and individuals.
Strong bias for hosts of a specific gender or age
class is evident in many water mite species with
terrestrial larvae, apparently resulting from differences
in the likelihood of exposure to mites of males and
females of various ages within host populations that
are determined by their behavior (Smith, 1999).
Preparasitic attendance of hosts by aquatic larvae prevents gender based behavioral differences in exposure
to parasitism and synchronizes initiation of larval engorgement with emergence of adult hosts. In these
mites, infestation rates of male and female hosts of a
given species at a particular time may differ in nature,
but partitioning the data by date usually removes any
apparent pattern. Gender-based selection of hosts
among colonizing mites was demonstrated by Lanciani
(1988). He found that larval Arrenurus exhibited
significant but small preference for female pupae of
Anopheles crucians, given equivalent exposure to both
genders of host pupae, and adjusting for any sexrelated differences in body size. Among mosquitoes,
dragonflies and some other host groups, the likelihood
of an adult parasitized by mite larvae returning to water is strongly influenced by its sex, and this should result in selective pressure for larval mites to recognize
the gender of potential hosts.
iv. Infecting host After encountering a potential
host, terrestrial larvae locate an appropriate attachment site and begin to feed. Species with larvae that exhibit preparasitic attendance must transfer from the
penultimate instar of the host to the imago during ecdysis. Larval Arrenurus cling passively to the preimaginal host instar, becoming active only when it begins to
molt. Interestingly, many larvae of species parasitizing
mosquitoes appear to have difficulty in locating the
split in the pupal exuvia and some that do find it are
unable to penetrate the surface film quickly enough to
reach the adult mosquito as it emerges. As a result, between 30 and 50% of preparasitic larvae fail to infect
the adult host (Smith and McIver, 1984b). Larvae of
Arrenurus species parasitic on odonates are carried out
of the water on the penultimate instar of the host and
almost all of them successfully move from the larval
exuvia onto the adult odonate (Mitchell, 1969b). Larvae of species of the genera Atractides, Hygrobates and
Unionicola pierce the pupal cuticle of their chironomid
hosts and embed their chelicerae in the pharate adult.
This behavior virtually ensures successful transfer to
the adult because the larvae are passively pulled
through the pupal exuvia as the host emerges (Böttger
1972b).
Little is known about the mechanisms by which
mite larvae select sites on hosts, and virtually all available information pertains to a few species of the genus
Arrenurus. Site selection of larval Arrenurus on
odonates appears to be largely a function of timing
(Mitchell, 1961). Progress of the host ecdysis at the
point when mite larvae become active determines
where the mites first come in contact with the host, and
they typically attach close to that point. Larvae experimentally removed from the host and placed elsewhere
on the body readily attached there. In contrast, there
appears to be little correlation between the process of
host ecdysis and site selection in larval Arrenurus parasitizing mosquitoes. For example, larval Arrenurus
kenki almost always attach between the head and thorax when infesting adult Aedes excrucians but to the
abdomen when attacking Aedes cinereus, without reference to the time taken by the host during ecdysis.
v. Attaching to host and engorgement Larval
mites attach to hosts by using their pedipalps to stabilize the body and produce the leverage needed to pierce
the cuticle of the host with their chelicerae. Larvae of
some species of Arrenurus secrete a substance which
solidifies and apparently helps to cement the mite to
the host (Åbro, 1984), but this has not been observed
in other cases (see Redmond and Lanciani, 1982).
After attachment, larvae of several genera (e.g.,
Hydrachna, Rhyncholimnochares, and Arrenurus) are
known to form a feeding tube, or stylostome, in the tissues of the host (see Smith, 1988), and these structures
are probably produced by larvae of species belonging
to many other genera. Gross structure of stylostomes
appears to be consistent for species within a genus, but
differs greatly among genera. For example, stylostomes
of larval Hydrachna (Davids, 1973) and Rhyncholimnochares are multibranched and open-ended, those of
larval Arrenurus are single blind tubes (Åbro, 1979;
Redmond and Hochberg, 1981) and the known example of a stylostome of a pionid appears to be a single,
open-ended tube (Gledhill, 1985). Details of stylostome
morphology appear to be constant for a given species
of mite regardless of host but differ among mite
species, leading to the conclusion that they are produced by larval mites rather than their hosts (Lanciani
and Smith, 1989). Stylostomes are acellular, consisting
of acid mucopolysaccharides, and have no perforations
other than a terminal pore, when present. The mechanism of stylostome function is not well understood.
16. Water Mites (Hydrachnida) and Other Arachnids
Davids (1973) argues that the stylostomes of larval Hydrachna are formed by coagulation of host hemolymph
in reaction to mite saliva, and that the mites imbibe hemolymph from the host. Stylostomes of Arrenurus larvae are typically surrounded by a fluid-filled abscess,
and it has been suggested that the mites feed upon liquefied tissues in much the same way as larval chiggers
do (Pflügfelder, 1970; Åbro, 1982, 1984). It is possible
that the stylostomes of larvae in these remotely related
clades are non-homologous.
Duration of the period of larval engorgement
varies considerably among water mite taxa. Larvae of
Limnochares aquatica and Hydryphantes tenuabilis require 6 – 13 days to engorge (Böttger, 1972a; Lanciani,
1971a; Smith, 1989), but those of various species of
Hydrachna and Eylais may spend from 2 weeks to 10
months on the host (e.g., Davids, 1973; Smith, 1977,
1988; Wiggins et al., 1980). Larvae of some species of
Eylais can complete engorgement within 2 weeks at the
time of year when female hosts are producing eggs, but
apparently diapause in a partially engorged state at
other times of the year (Smith, 1988). Presumably,
growth rate of larvae in these groups is generally dependent on physiological condition of the host. Larvae
of more recently evolved groups associated with shortlived insects, especially nematocerous Diptera, require
shorter times for engorgement. Those of pionid species
inhabiting temporary pools, such as Piona napio and
Tiphys americanus, can engorge in as little as 24 h on
their chironomid hosts. Larval Arrenurus feeding on
mosquitoes take approximately 6 days for full engorgement (Mullen, 1974).
Larval growth during engorgement is also highly
variable. Larvae parasitic on nematocerous flies increase only modestly in size. For example, larvae of
Limnesia maculata and Unionicola crassipes expand to
between 3 and 4.3 times their original volume on their
chironomid host and those of Arrenurus species feeding
on mosquitoes increase in volume about 16-fold
(Münchberg, 1938; Böttger, 1972b). Larvae of other
species of Arrenurus parasitic on odonates augment
their volume 80 to 90 times, and larval Hydrachna and
Eylais grow substantially, increasing up to 600-fold in
size in some cases (Davids, 1973; Fig. 427).
Larvae parasitizing long-lived hosts for extended
periods risk premature detachment when the host
molts. Caste exuvia of nepid and belastomatid bugs are
commonly found with larval and protonymphal Hydrachna still attached. Larvae of Limnochares aquatica
retain their mobility and can transfer from one host instar to the next to continue engorgement.
vi. Detaching from host Following engorgement,
mite larvae must detach from the host and return to an
565
aquatic habitat suitable for postlarval development.
Larvae of various taxa may use host behavior, along
with mechanical, visual, and chemical stimuli, as cues
to initiate detachment (Böttger, 1972a, b). Environmental cues (such as moisture) may be sufficient to induce
detachment in some species, but physiological cues
from the host are essential in others (Smith and Laughland, 1990). Larvae of the Eurasian species Arrenurus
cuspidator parasitic on the damselfly, Coenagrion
puella require two simultaneous cues to stimulate detachment, high humidity and either mating or oviposition behavior by the host (Rolff and Martens, 1997).
Presumably, the mite larvae can detect hormones or
some other physiological correlate with reproductive
behavior in the host.
Engorged larval mites risk detachment from hosts
into unsuitable habitats. Fully fed larvae with limited
mobility falling on dry ground quickly perish, and
those introduced into unsuitable aquatic habitats may
fail to survive or reproduce. At least some species of
Arrenurus parasitic on odonates are regularly introduced into lakes that apparently cannot support their
reproduction (Mitchell, 1964a, 1969b). A large proportion of Cenocorixa bifida waterboatmen living in
inland saline lakes of central British Columbia are parasitized by nonresident species of Eylais and Hydrachna by late autumn (Smith, 1977).
Fully fed larvae are capable of only limited movement, and those successfully re-entering water typically
seek out plant material or detritus, attach themselves
by their chelicerae and become quiescent. Engorged larvae of Piona and Arrenurus may attempt to swim or
crawl for up to 2 days before transforming to
protonymphs .
3. Protonymph (Nymphochrysalis)
Engorged larvae of all groups of water mites enter
a quiescent protonymph stage during which larval tissues are resorbed and reorganized and the deutonymph
develops. After a few days fully formed deutonymphs
emerge from the larval skins and become active.
Mites which parasitize large, long-lived insects, including Hydrachna, most Eylaoidea, and certain
Hydryphantidae, apotypically pass through the
protonymphal stage while still attached to the host,
within the larval integument. This trait is correlated
with extreme growth during the parasitic phase which
renders engorged larvae incapable of locomotion.
Along with other adaptations, extended attachment to
the host allows many of these mites to exploit periodically temporary habitats efficiently.
Members of Arrenurus planus Marshall and A.
ventropetiolatus Lavers survive for up to 10 months of
the year in the dry basins of vernal temporary pools in
566
Ian M. Smith, David R. Cook, Bruce P. Smith
eastern and western North America, respectively, as diapausing protonymphs and pharate deutonymphs.
These mites require a period of desiccation, and probably cold temperatures, to release them from diapause
(Münchberg, 1937; Wiggins et al., 1980).
4. Deutonymph
In all known species the deutonymphal instar is active and predaceous and resembles the adult, but is sexually immature. Deutonymphs typically exhibit relatively undeveloped idiosomal sclerites and chaetotaxy
compared to adults. They have only a rudimentary (or
provisional) genital field bearing an incomplete complement of acetabula. They feed voraciously, often preying
upon immature instars of the same taxa of insects they
parasitized as larvae (see Table I). The deutonymphal
stage varies in duration from a few days or weeks in
early derivative groups such as Hydrachnidae, Eylaoidea and certain Hydryphantidae, to several months
in many groups of Lebertioidea, Hygrobatoidea and
Arrenuroidea. Deutonymphs of some hygrobatoid families are especially long-lived. Those of Pionidae typically live for many months through summer and the
following winter, and this trait has pre-adapted certain
species to exploit annually temporary pools. In these
pionids, deutonymphs are able to endure the dry phase
of the cycle in damp retreats in the substrate (Smith,
1976a; Wiggins et al., 1980).
The deutonymph is the primary growth instar in
most groups of water mites, and body size increases
dramatically during this stage. On attaining adult size
deutonymphs prepare to transform to tritonymphs by
embedding their chelicerae in plant tissue or soft detritus and becoming inactive.
5. Tritonymph (Imagochrysalis)
During this stage further structural reorganization
occurs to produce the adult. This final metamorphosis
is rapid, and adults typically emerge within a few days.
Mature, fully fed deutonymphs anchor themselves
to living or decaying plant material using the legs and
mouthparts as they approach transformation to the
tritonymph stage and become quiescent. Tritonymphs
of several species of Pionidae inhabiting vernal temporary pools are regularly found attached to aquatic
mosses in clusters, with each mite attached at a leaf
axil. Conspicuous clumps of up to 500 tritonymphs
comprising several species of the genera Tiphys and Piona can be observed in moss mats at the edges of these
pools as the ice is melting in early spring. The significance of this clustering is not known. The tritonymphal
instar remains enclosed by the deutonymphal exoskeleton, and the pharate adult may often be seen forming
within it. Mites of various species of Arrenurus typi-
cally spend 4 to 5 days in the tritonymph stage under
laboratory conditions at 20°C. Water mites appear to
be particularly vulnerable to poor water quality during
the tritonymph stage, perhaps because the metamorphosing adults must respire by diffusion across three
levels of cuticle. For example, when dead and decaying
prey are present in laboratory cultures, tritonymphs of
Arrenurus may take up to 17 days to complete development and mortality rates of this instar may exceed
50%, despite minimal mortality among deutonymphs
and adults living in the same container.
6. Adult
a. Sclerotization of Exoskeleton Adults emerge
from the deutonymphal integument in teneral condition with soft, pliable, and colorless sclerites. They immediately become active, crawling or swimming about
while sclerotization of the body is completed and distinctive color patterns become evident. Adults of the
various groups display a wide range of body shapes
and arrangements of idiosomal sclerites, primarily as
adaptations for living in aquatic habitats of varying
depth, turbulence, temperature, and substrate type.
Shortly after emergence, both males and females mature sexually and begin to mate. In many taxa, males
tend to emerge and become mature a few days earlier
than conspecific females, and are ready to mate as soon
as females become active in the habitat. Adults of certain Limnochares (Böttger, 1972a), and possibly some
Arrenurus (Imamura, 1952), undergo supernumerary
molts.
b. Mating and Spermatophore Transfer The adult is
a highly specialized reproductive instar in water mites,
as clearly evidenced by the remarkable behavioral modifications, and associated morphological adaptations,
that have developed in various groups to facilitate successful spermatophore transfer (Smith and Oliver,
1986; Proctor, 1992a). Water mites exhibit a greater
variety of spermatophore transfer methods than any
other group of arachnids (Proctor, 1991b). Water mite
genera can be grouped into categories based on degree
of interaction between males and females during spermatophore transfer (Proctor, 1992b).
i. Dissociation during transfer The plesiotypical
behavior pattern in water mites, complete dissociation
involving no chemical communication or physical contact between the sexes, has been observed in species of
a wide variety of early derivative genera (see Proctor,
1992a). In these species, males deposit stalked spermatophores on the substrate for subsequent retrieval
by conspecific females, and those of some species have
been observed to deposit large numbers when isolated
16. Water Mites (Hydrachnida) and Other Arachnids
in small containers of water. After finding the spermatophores, perhaps using pheromonal cues, females
pick them up in the gonopore. An apotypical variant of
this, incomplete dissociation requiring chemical or
physical cues from females to that stimulate males for
depositing spermatophores, occurs in several unrelated
genera (Proctor, 1992a). Dissociation during spermatophore transfer is widespread in actinedid mites
and probably occurs in most groups of water mites
with unmodified males.
ii. Association during transfer A more interactive
type of mating, pairing of males and females for indirect spermatophore transfer, has been observed in several species (see Proctor, 1992b). In these cases, males
deposit spermatophores on the substrate, then actively
assist females to find them and pick them up. Males of
some Eylais lead receptive females in a circular dance,
stopping repeatedly to deposit spermatophores at the
same location on the circumference of the route.
Halfway through each revolution of the dance, the female pauses with the male over his mass of spermatophores and picks up one or more of them in her
gonopore (Lanciani, 1972). Males of Neumania papillator apparently exploit the hunting behavior of females to attract them to spermatophores (Proctor,
1991a, 1992a, c). Upon encountering a female, males
vibrate their legs rapidly, simulating the stimuli produced by copepod prey, until she responds with characteristic ambush posture and aggression. The males then
turn away, lay down a spermatophore net and rapidly
fan water to create a current over the net toward the
female, presumably directing pheromones toward her.
A receptive female then ceases aggression, approaches
the spermatophore net, and begins to collect spermatophores. Males of some species of Unionicola also
use leg-trembling to generate vibrational cues to attract
females to spermatophores (Proctor, 1992a, c).
iii. Copulation during transfer Pairing with direct spermatophore transfer has evolved independently
in many water mite lineages. Active transfer of spermatophores to females is accomplished either by
modified appendages or idiosomal sclerites of males or
by direct gonopore-to-gonopore contact. Proctor
(1992b), following Hinton (1964), considers both of
these options as examples of copulation.
In Unionicola intermedia, a species that inhabits
the mantle cavity of freshwater mussels, males retrieve
their own spermatophores and carry them between the
genua or tibiae of their hind legs while seeking receptive females. On finding a female, the male crawls beneath her and places his spermatophores in her gonopore (Hevers, 1978). Males of many groups of
567
Hygrobatoidea produce unstalked spermatophores
which they carry between modified claws on the tarsi
of their flexed third pair of legs before transferring
them directly into the gonopore of the female. This
type of mating has been observed in various species of
Feltria (Motas, 1928), Forelia (Uchida, 1940), Tiphys
(Viets, 1914; Mitchell, 1957c), Piona (Koenike, 1891;
Mitchell, 1957c; Böttger, 1962; Smith, 1976a) and
Brachypoda (Motas, 1928; Halik, 1955). In each case,
the male uses specially modified appendages, usually
the fourth pair of legs, to hold the female in a characteristic posture so that he can transfer spermatophores
to her gonopore by simply extending his third pair of
legs. This mating behavior probably occurs in all water
mites whose males have highly modified leg segments
but no elaborate caudal development.
A variety of complex modes of direct spermatophore transfer occur in the genus Arrenurus.
Males of the various species have the idiosoma modified posteriorly to form a characteristically shaped
cauda (Figs. 419, 420). A receptive female mounts a
male, positioning herself so that her gonopore rests
over the back edge of his cauda. Secretions from glandularia of the male temporarily cement the pair together, and the male then deposits one or more stalked
spermatophores on the substrate. In some species, the
male then rocks his body down and forward to bring
the gonopore of the female in contact with the spermatophore (Lundblad, 1929; Böttger and Schaller,
1961; Böttger, 1962). In others, the male uses his petiole, a cup-shaped posterior appendage of the cauda, to
scoop up spermatophores into masses which he then
inserts into the gonopore of the female (Böttger, 1965).
Variations of this behavioral pattern occur throughout
the many species of Arrenurus, along with the multitude of different modifications in caudal morphology
that characterize the males. This type of mating probably can be inferred in all groups of water mites with
elaborate caudal development in males.
Finally, direct transfer of spermatophores from
gonopore to gonopore during copulation occurs in certain species of Eylais (Böttger, 1962; Lanciani, 1972)
and in the arrenuroid genera Midea (Lundblad, 1929)
and Nudomideopsis (I.M. Smith, unpublished data). In
some of these cases, males either have the gonopore
protruding from the venter on a cone-shaped projection (Eylais) or are equipped with winglike appendages flanking the gonopore (Midea) to facilitate
transfer.
Clearly, copulation during spermatophore transfer
has evolved many times from antecedent conditions involving association with complex sexual interaction. As
defined by Proctor (1991b), copulation has developed
independently no fewer than 91 times within the water
568
Ian M. Smith, David R. Cook, Bruce P. Smith
mites, including at least three times within a single family, Pionidae (Mitchell, 1957c; Smith, 1976a). Direct
transfer from gonopore to gonopore certainly developed independently in Eylais and Arrenuroidea.
The often profound modification of the male idiosoma and appendages that is associated with diverse
modes of spermatophore transfer emphasizes the highly
specialized mating function of the adult instar in water
mites. Males in many groups mate and die within a few
days of emerging. Those of some groups, such as Arrenurus, may live for several weeks. Males of some
genera of Foreliinae and Tiphyinae are so modified
morphologically that they are unable to move about
easily, and are restricted to crawling slowly over the
substrate using the anterior pairs of legs.
Sex pheromones appear to mediate pairing in at
least some species of water mites. Males of Limnesia
undulata increase spermatophore deposition when females are present, and produce comparable numbers
when placed in containers from which females have recently been removed (Proctor, 1992a). The vigorous
fanning of water over spermatophore nets toward females by males of Neumania papillator strongly suggests chemical communication (Proctor, 1991a). Females of various species of Arrenurus produce
male-arrestant / attractant pheromones that can be extracted from water (B.P. Smith and Hagman, unpublished data). Unlike insect pheromones which are often
species-specific, the pheromones of Arrenurus species
often elicit response in several closely related and cooccurrent species (B.P. Smith and Florentino, unpublished data). There is also evidence that males of some
species of Arrenurus produce pheromones attractive to
females (Baker, 1996).
Males appear to be much less common than females in many genera of water mites (Meyer and
Schwoerbel, 1981; Proctor, 1997). This may in part be
due to the shorter life span of males. Biased sex ratios
can also result from protandry which has been demonstrated in at least some water mite species (Proctor,
1989, 1992b). Females of Arrenurus manubriator produce strongly gender-biased clutches (Proctor, 1997;
B.P. Smith, unpublished data). The bias for males or females appears to be consistent among successive
clutches of eggs for a given female and appears to be a
trait largely inherited from the maternal parent (B.P.
Smith, unpublished data). Water mites appear to be
diplo-diploid, with no recognizable sex chromosomes
(Sokolow, 1954).
Mated females may live for many months, continuing to feed while they produce several clutches of eggs.
In many groups, mating occurs in late summer and fertilization is delayed while females overwinter. These females lay their eggs after they have been fertilized early
in the following spring or early summer. In contrast, females of species inhabiting temporary pools must lay
their eggs within a few days of mating to ensure that
their offspring reach a life-history stage capable of
avoiding or withstanding the dry period before water
disappears from the habitat (Smith, 1976a; Wiggins
et al., 1980).
Preliminary evidence suggests that the first male to
mate with a female has almost total sperm precedence
over subsequent mates. Given that sperm is regularly
stored for many months over winter, there must be substantial selective pressure for behavioral and phenological adaptations to gain early access to females. Guarding of tritonymphs (presumably females) by young
males has been observed in Arrenurus planus.
Males do not seem to discriminate between virginal and previously mated females. Males of Neumania papillator attempt to mate with all females they
encounter. They eventually produce significantly
greater numbers of spermatophores for virginal females, but this appears to be the result of avoidance
behavior by inseminated females rather than discrimination by males (Proctor, 1991a). Males of Arrenurus
manubriator choose larger females over smaller ones,
perhaps selecting for greater egg production capacity
(Proctor and Smith, 1994; Smith and Bovenzi, unpublished data).
D. Habitats and Communities
Throughout water mite evolution, successful invasion and exploitation of new habitats has depended on
the development of compatible adaptive strategies for
the various instars. Larval traits tend to promote parasitism and dispersal on hosts, while those of deutonymphs and adults favor feeding, growth and reproduction in water. Water mite species are regularly
provided with opportunities to colonize new habitats
by the passive transport of their larvae on hosts, and
this has been a primary mechanism promoting speciation and divergence over time. Nevertheless, in modern
communities most species and many monophyletic
groups representing genus or family level taxa are restricted to one or a few similar types of habitats. The
strong correlation between certain clades and habitats
suggests that conservative factors such as adaptive requirements for locating hosts, prey, mates, and oviposition sites tend to constrain adaptive radiation. It also
indicates that successful invasion of new habitats usually resulted in the rapid evolution of significant new
life-history, behavioral and morphological traits.
The communities living in the major types of freshwater habitats in North America are outlined below
and in Table I.
16. Water Mites (Hydrachnida) and Other Arachnids
1. Springs (Including Seepage Areas)
Species of 47 genera live in moss and wet detritus
associated with rheocrenes and helocrenes. Members of
ancient clades that may well have originated in this
type of habitat are soft-bodied (e.g., Tartarothyas), or
partly to fully sclerotized (e.g., Thyas, Panisopsis).
Adults of derivative groups that apparently invaded
spring and seepage habitats from flowing water more
recently have entire dorsal and ventral shields
(e.g., Paramideopsis — Fig. 425, certain Mideopsis,
Laversia). Most spring inhabiting species are coldadapted stenophiles, but some Wandesia (subgenus
Partnuniella) and all Thermacarus live only in hot
springs. Over 115 species in 57 genera and 25 families
have been identified from spring habitats in Canada including interstitial taxa (Smith, 1991a).
2. Riffle Habitats
Species of 49 genera live on the substrate in rapidly
flowing areas of streams and rivers. Diverse communities from a single location may contain up to 50 species
in 30 genera. Members of a few early derivative taxa
are soft-bodied (e.g., Protzia), but adults of most groups
are both strongly flattened and well sclerotized (e.g.,
Kongsbergia — Fig. 421, Aturus — Fig. 422). Most
rheophilic species are cold- or cool-adapted stenophiles.
3. Interstitial Habitats
Species of 55 genera live in sand and gravel deposits to depths of 1 m or more, mainly in the hyporheic zone of streams and rheocrenes. Deutonymphs
and adults crawl through the spaces between particles,
often with considerable speed and agility. Members of
most early derivative taxa are soft-bodied (e.g., Rhyncholimnochares) and some are also vermiform (e.g.,
Wandesia). In adults of more recently derived groups,
the dorsum and venter are partly covered by various
plates or small shields (e.g., Clathrosperchon, Omartacarus, Atractides), or are well sclerotized with either
entire dorsal and ventral shields or greatly expanded
coxal plates (e.g., Frontipodopsis, Aturus — Fig. 422,
Uchidastygacarus — Fig. 424, Arenohydracarus). Mites
adapted to interstitial habitats have reduced eyes and
lack pigmentation of the integument.
4. Stenothermic Pools
Species of 47 genera live in silty substrates in
spring-fed pools, fen pools and depositional areas of
streams. Members of ancient taxa are soft-bodied (e.g.,
Pseudohydryphantes), while adults of early derivative
clades exhibit various degrees of sclerotization ranging
from slight enlargement of dorsal and ventral plates
(e.g., Teutonia, Wettina, Forelia) to development of en-
569
tire dorsal and ventral shields (e.g., Momonia, Geayia).
Adults of certain taxa which invaded pools relatively
recently from lotic and hyporheic habitats have retained dorsal and ventral shields, though they have become secondarily adapted for swimming (e.g., Axonopsis, Mideopsis). Certain groups inhabiting depositional
areas of streams (e.g., Teutonia, Oxus, Wettina) include the fastest and most agile swimmers among the
water mites. Most inhabitants of pools are cooladapted stenophiles.
5. Lakes (Including Permanent Ponds, Marshes,
Swamps and Bogs)
Species of 39 genera inhabit large bodies of standing water. Members of ancient and early derivative genera are generally soft-bodied (e.g., Hydrodroma, Limnesia, Hygrobates, Huitfeldtia). Adults of many genera
in more recently evolved clades have extensive dorsal
and ventral shields (e.g., Koenikea — Fig. 423), in at
least some cases because the group has secondarily invaded standing water from lotic or hyporheic habitats
(e.g., Axonopsis, Mideopsis). Most species are strong
swimmers and have fringes of long, slender setae on the
genua and tibiae of the second, third, and fourth pairs
of legs (Fig. 290). These swimming setae help to propel
the mites as they move about at the surface of the
substrate or among plants. Some species of Unionicola
and Piona have especially long swimming setae, and
are essentially pelagic or planktonic (Riessen, 1982). A
few species which lack swimming setae either walk or
crawl actively on plants and detritus, reflecting recent
rheobiontic ancestry (e.g., Sperchon, Torrenticola, certain Mideopsis, Neoacarus). Members of Tyrrellia are
unusual in that they inhabit wet litter at the edges of
permanent bodies of water, including lakes, where they
crawl about at the surface film. Many species of Unionicola and the only known species of Najadicola are obligate parasites or commensals in the mantle cavity of
molluscs in both lentic and lotic habitats (Mitchell,
1955; Simmons and Smith, 1984; Gledhill, 1985;
Vidrine, 1986).
More than 25 species in each of the remarkable
genera Arrenurus and Piona often occur together in a
single small lake. Mitchell (1964a, 1969a) concluded
that behavioral flexibility exhibited by larval Arrenurus
in selecting and exploiting their odonate hosts is the
main factor permitting such high levels of sympatry,
and that competition between deutonymphs and adults
of the various species is probably not significant in natural communities.
Most limnophilic taxa tolerate a broad range of
temperatures which they encounter in weed beds and
silty substrates in the littoral and sublittoral zones.
However, a few are cold-adapted stenophiles restricted
570
Ian M. Smith, David R. Cook, Bruce P. Smith
to either arctic-alpine glacial lakes (e.g., Acalyptonotus)
or the profundal zone of oligotrophic lakes (e.g., Huitfeldtia, Laversia). Certain species of some genera (e.g.,
Limnochares, Hydrodroma, Limnesia, Piona and Arrenurus) are highly tolerant of extreme thermal and
chemical regimes, and are able to inhabit desert
sloughs, alkaline lakes or acid bogs. The water mite
fauna of wetland habitats in Canada was treated by
I.M. Smith (1987).
6. Temporary Pools
Species of 13 genera live in annually temporary
pools. Members of ancient genera that may have originated in this type of habitat range from soft-bodied
walkers (e.g., Thyas) and swimmers (e.g., Eylais) to
mites with unique arrangements of integumental plates
that either crawl (e.g., Piersigia) or swim awkwardly
(e.g., Hydrachna, Hydryphantes). Species of derivative
groups that apparently invaded temporary pools secondarily tend to be relatively good swimmers (eg:Tiphys,
Piona, Arrenurus). Water mites inhabiting temporary
pools are adapted either to avoid (e.g., species of Hydrachna and Eylais) or to endure the dry phase of the
annual cycle (Wiggins et al., 1980).
E. Ecology
The parasitic larvae and predaceous deutonymphs
and adults of water mites have direct and almost certainly significant effects on the size and structure of insect populations in many habitats (Smith, 1983, 1988;
Lanciani, 1983; Proctor and Pritchard, 1989). Unfortunately, their impact has rarely been measured accurately because of the routine neglect of mites in ecological studies of freshwater communities. Due to their
small size and often cryptic habits, mites are usually
absent or seriously under-represented in samples that
are collected using standard techniques for capturing
insects and crustaceans. Most entomologists are unfamiliar with mites, and tend either to disregard them as
too poorly known ecologically and difficult taxonomically or to lump them together in a meaningless way,
when conducting community studies. Failure to include
realistic assessment of the roles played by water mites
often results in seriously flawed analyses of the structure and dynamics of freshwater communities.
1. Impact as Parasites
a. Effect on Individual Hosts The impact of larval
water mites on host insects was reviewed by Lanciani
(1983) and B.P. Smith (1988). Numerous laboratory
studies have demonstrated reduced survival or
longevity of various insect hosts parasitized by larval
water mites, including mosquitoes (Lanciani and Boyt,
1977; Lanciani, 1987), ceratopogonid midges (Lanciani, 1986a), chironomid midges (Weiberg and Edwards, 1997), juvenile water striders (Smith, 1989),
backswimmers (Lanciani, 1982) and damselflies
(Forbes and Baker, 1990, 1991; Leung and Forbes,
1997). Lanciani (1983) concluded that the ratio of larval mite weight-to-host weight is a good indicator of
the probable impact of parasitism on the host. Studies
of parasite-induced mortality in field populations have
not been conclusive (Robinson, 1983; Åbro, 1990a;
Smith, 1999). There is evidence of occasional crashes
of host populations related to parasitism by mites
(Smith, 1988).
Water mite parasitism reduces egg production of
insect hosts. The most pronounced effects have been reported for Hemiptera infested with larval Hydrachna
and Eylais (Davids, 1973; Smith, 1977), but parasitized nematocerous flies also show lowered fecundity
(Smith and McIver, 1984c; Lanciani and Boyt, 1977).
Development of juvenile instars is retarded in backswimmers (Notonectidae) parasitized by larvae of Hydrachna virella (Lanciani and May, 1982) and in water
striders (Gerridae) attacked by Limnochares aquatica
(Smith, 1989).
Mite parasitism influences frequency of foraging,
intensity of territorial behavior and likelihood of mating in males of the damselfly Enallagma ebrium
(Forbes, 1991). Mite parasitism apparently reduces the
capacity of the damselfly Nehalennia speciosa to fly
and disperse (Reinhardt, 1996). The corixid Cymatia
bonsdorfi increases feeding activity when parasitized by
larvae of Hydrachna conjecta (Reilly and McCarthy,
1991).
Some hosts exhibit strong avoidance behavior in
the presence of larval water mites, involving grooming
behavior to dislodge them. Larval Ischnura verticalis
typically respond to fish by reducing movement, but in
the presence of both fish and preparasitic larvae of Arrenurus pseudosuperior exhibit unrestrained grooming
and mite-avoidance behavior despite the increased risk
of detection by predators (Baker and Smith, 1997).
b. Effect on Population Structure of Hosts Larval
water mites regularly parasitize 20 – 50% of adults in
natural populations of aquatic insects in such diverse
families as Corixidae (Hemiptera), Dytiscidae
(Coleoptera), Libellulidae (Odonata), Culicidae, and
Chironomidae (Diptera). Almost all individuals are
parasitized in some host populations, and some may
carry remarkably high mite loads. We have seen examples of over 1000 larvae of Arrenurus parasitizing a
dragonfly (Erythemus simplicicolis), over 350 larvae of
Limnochares aquatica on a water strider (Gerris comatus), over 50 larvae of Arrenurus on a ceratopogonid
16. Water Mites (Hydrachnida) and Other Arachnids
midge (Bezzia sp.) and over 40 larvae of Arrenurus
danbyensis on a mosquito (Coquillettidia perturbans).
Individual chironomid midges emerging from
mesotrophic lakes in the Great Lakes Basin often carry
20 or more mite larvae representing as many as seven
different genera.
The distribution of parasitic larval water mites typically follows a clustered pattern within a host population, with a few host individuals heavily infested and
most carrying few or no mites (e.g., Smith and McIver,
1984a; Smith, 1988). Under natural conditions, clustering may reflect differences in the temporal and spatial
distributions of host-seeking larval mites and their potential hosts. However, clustering also occurs in laboratory experiments involving synchronous exposure of
insects to mites, where duration of exposure and spatial heterogeneity are controlled. This suggests either
initial differential susceptibility to parasitism among
host individuals or facilitation of additional loading on
hosts that have become parasitized.
Intensity of infestation by larval Arrenurus appears
to be negatively correlated with body size and condition (i.e., ratio of mass to body size) of their damselfly
hosts in at least some cases (Forbes and Baker, 1991;
Leung and Forbes, 1997). This may be a result of more
vigorous avoidance behavior by larger and more robust
potential hosts (Smith and McIver, 1984b; Forbes and
Baker, 1990). Larval Arrenurus have greater intensity
of parasitism on damselflies with asymmetry in
forewing length, but it is not clear whether increased
infestation is a response to or a cause of this asymmetry (Bonn et al., 1996).
Behavioral subtleties can increase vulnerability of
hosts of certain gender or age classes to colonizing larval mites (Smith and Cook, 1991), with resulting differentials in parasite loads (Smith, 1988, 1999). Observed differences in parasite loads among sex and age
classes in natural populations can prove useful for inferring the behavior of the host and parasite and the
age structure of the host population (Smith, 1999).
c. Effect on Other Mites Sharing the Same Host
Clustering of parasitic larvae on certain host individuals results in intraspecific competition between mites on
those hosts that can affect their growth and survival
(Lanciani, 1976, 1984, 1986b), and leaves many hosts
underutilised. Consequently, clustering apparently both
invokes density-dependent control of mite populations
and keeps them below carrying capacity, effectively
precluding interspecific competition.
Parasitism of a host individual by larvae of more
than one species of water mite occurs more commonly
than would be expected by chance (Stechmann, 1980;
Smith and McIver, 1984a; Lanciani and McLaughlin,
571
1989). In addition, many species of water mites exploit
a variety of related host species. As a result, parasitic
associations often involve complex assemblages with
considerable overlap in exploitation of host resources,
including use of similar attachment sites (Kouwets and
Davids, 1984). Interspecific competition appears to be
negligible in these assemblages. Consequently, assuming that larval success is the limiting factor determining
population size in water mite species, there is probably
little competition for food resources in communities of
deutonymphs and adults. This may help to explain the
remarkably high levels of species richness in many water mite communities.
2. Impact as Predators
The feeding habits of deutonymphal and adult water mites were reviewed by Böttger (1970), Gledhill
(1985) and Proctor and Pritchard (1989). Data for
species inhabiting Canadian wetlands were reviewed by
I.M. Smith (1987). Deutonymphs and adults of free-living species are voracious predators on a wide range of
small aquatic organisms. Some species at least occasionally scavenge on recently killed organisms. Most
species exhibit marked preferences for prey of a particular size with particular morphological and behavioral
attributes. Species of Hydrachna, Thyas, Hydryphantes
and Hydrodroma are specialized predators of arthropod eggs (Davids, 1973; Mullen, 1975; Lanciani, 1978,
1980; Wiles, 1982; Meyer, 1985). Many members of
Eylais and Piersigia and the large genus Arrenurus appear to depend on ostracod crustaceans as prey (although some Eylais and Arrenurus feed on cladocerans), adults of Neumania ambush copepods, pelagic
and planktonic species of Unionicola and Piona are
highly efficient predators of cladoceran crustaceans and
species of many genera of Lebertioidea, Hygrobatoidea
and Arrenuroidea prey upon larval Chironomidae
(Böttger, 1970; Proctor, 1991b; Proctor and Pritchard,
1989, 1990; also see Table I). Some species of Limnesia
and Piona appear to be more opportunistic in their
choice of prey, and frequently even attack other mites
in crowded containers (Elton, 1922). Parasitism of
freshwater molluscs during the deutonymphal and
adult stages has evolved at least twice in the superfamily Hygrobatoidea. Many species of Unionicola inhabit
the mantle cavity of mussels and snails (Mitchell, 1955;
Vidrine, 1986) and the pionid Najadicola ingens has
similar habits (Simmons and Smith, 1984).
Deutonymphs and adults frequently prey upon the
immature instars of the same species of insects that
they parasitize as larvae (Davids, 1973; Wiles, 1982;
Proctor and Pritchard, 1989). Preliminary indications
suggest that many species of mites are specialized to exploit one group of insects throughout their life history,
572
Ian M. Smith, David R. Cook, Bruce P. Smith
a strategy that would presumably increase the probability of both spatial and temporal co-occurrence of the
mites and their preferred hosts and prey.
Some water mites exhibit feeding rates in experimental studies that are high enough to be considered as
potentially important influences on the size and structure of prey populations in nature (Paterson, 1970;
Mullen, 1975; Lanciani, 1979; Wiles, 1982). Gliwicz
and Biesiadka (1975) reported that individuals of Piona limnetica consume 10 – 20 cladocerans per day in a
neotropical lake, and estimated that members of this
species could reduce the average standing crop of prey
by 50% each week. Adults of Hygrobates nigromaculatus, with reported population densities as high as 1000
individuals / m2, apparently consumed an average of
14,500 larvae of the chironomid Cladotanytarsus mancus per square meter for two consecutive years in a European lake, accounting for a 50% mortality rate in the
prey population (Ten Winkel et al., 1989).
Tactile cues appear to be of primary importance to
predatory water mites (Böttger, 1970; Davids et al.,
1981). Hunting mites often ignore stationary potential
prey even at close range, but immediately attack and
kill individuals of the same prey species after contact is
made. Adults of species of Piona typically swim in wide
arcs until colliding with potential prey. Upon contact,
the mites attack aggressively, and if initially unsuccessful, swim in tight spirals for several seconds until the
prey is encountered again (B.P. Smith, personal observation). Several species of Neumania and Unionicola
orient toward vibrations in the water as a cue to locate
potential prey (Proctor and Pritchard, 1990).
Water mites also use various cues to promote efficient prey capture. Adults of Hydrodroma despiciens
seem to respond to chemicals in the gelatinous coating
of the insect eggs on which they feed (Wiles, 1982).
Adults of a number of species show behavioral responses to chemical cues from prey organisms, and
Baker (1996) has demonstrated how the mites detect
these stimuli. Adults of both free living and parasitic
species of Unionicola react to host extracts suggesting
that they use chemical cues to initiate regional searching (Proctor and Pritchard, 1989; Roberts et al., 1978;
Dimock and LaRochelle, 1980; LaRochelle and Dimock, 1981).
3. Importance as Prey
Water mites are typically under represented in the
stomach contents of predatory freshwater vertebrates
(Modlin and Gannon, 1973; Pieczynski, 1976; Eriksson et al., 1980). Deutonymphs and adults at least occasionally form a significant part of the diet of fish and
turtles (Marshall, 1933, 1940; Knight, 1989), and we
have seen examples of stomach contents from individ-
ual brook trout and coregonids that consisted exclusively of several hundred adults of one species of Piona
and Hygrobates, respectively. It appears that fish can
develop narrowly focused search images for water
mites in exceptional cases.
Adults of many species of water mites eject viscous, sticky fluid from the glandularia when handled
roughly, apparently as a deterrent to predation. Experiments have shown that red-colored water mites are distasteful to fish which quickly learn to reject them as potential prey (Elton, 1922; Kerfoot, 1982). Kerfoot
(1982) concluded that red pigmentation in water mites
is aposematic coloration. However, most red water
mites live in temporary pools and seepage areas where
fish are absent. Terrestrial anystoid and parasitengone
mites, and members of many early derivative water
mite taxa, are predominantly red or orange suggesting
that suffusion of the body with red carotenoid pigments is plesiotypical for the entire lineage. This type
of pigmentation may have initially evolved in these
mites as protection from ultraviolet wavelengths in
sunlight (Garga, 1996). However, it is noteworthy that
few members of the derivative water mite superfamilies
Lebertioidea, Hygrobatoidea of Arrenuroidea have retained the plesiotypical condition. Most species in these
groups are not red, and the relatively small number
that are have acquired this coloration secondarily, as
explained above. Interestingly, most red species in these
groups also inhabit vernal temporary pools or springs.
Few studies have examined invertebrate predation
on water mites. Elton (1922) reported an instance of
predation by an adult dytiscid beetle on a species of
Hydrachna. Two authors have reported positive associations between fish and the mite Piona carnea and suggested both that fish prefer insects to mites as prey and
that predaceous insects, including hemipterans, larval
odonates and larval chaoborid midges, feed on the
mites (Eriksson et al., 1980; Punčochér and Hrbáček
1991).
4. Potential as Indicators of Environmental Quality
Species of water mites are specialized to exploit
narrow ranges of physical and chemical regimes, as
well as the particular biological attributes (including
physico-chemical constraints) of the organisms they
parasitize and prey upon. Consequently, water mites
should be exceptionally sensitive indicators of habitat
conditions and the impact of environmental changes on
freshwater communities. Preliminary studies of
physico-chemical and pollution ecology of the relatively well-known fauna of Europe have demonstrated
that water mites are excellent indicators of habitat
quality (Schwoerbel, 1959; Pieczynski, 1976; Biesiadka, 1979; Kowalik and Biesiadka, 1981; Kowalik,
16. Water Mites (Hydrachnida) and Other Arachnids
1981; Bagge and Merilainen, 1985; Cicolani and di
Sabatino, 1991; Steenbergen, 1993). Gerecke and
Schwoerbel (1991) analyzed changes in water mite
communities in the Danube River between 1959 and
1984 and concluded that they were highly correlated
with increased levels of organic pollution. One of the
species they studied, Hygrobates fluviatilis, appears to
be relatively resistant to pollution and increased in
dominance during the period of declining water quality. The results of these studies, along with our own observations in sampling a wide variety of habitats in
North America and elsewhere, lead us to conclude that
water mite diversity is dramatically reduced in habitats
that have been degraded by chemical pollution or physical disturbance.
There have been few laboratory studies of the tolerances of water mites to environmental variables.
Deutonymphs of Arrenurus manubriator have proven
to be more susceptible than other life-history stages to
iron in solution, probably resulting from the major
contribution of this instar to growth of the mites
(Rousch et al., 1997). Experiments to assess differential
susceptibility among life-history instars may provide
valuable insight into the mechanisms responsible for
observed changes in species populations in stressed
habitats.
Progress toward the establishment of essential
baseline documentation on water mite communities in
North America, however, continues to be hampered by
lack of sufficient systematic and ecological information
at the species level, and by inadequate sampling and
extraction procedures. Application of potentially useful information on water mites in assessing and monitoring the well-being of freshwater habitats and
communities demands clearer understanding of the individual niche requirements and trophic relationships
of species. This can be achieved by improving systematic knowledge of the group, and by developing more
appropriate collecting, observing, and rearing techniques.
F. Collecting, Rearing, and Preparation for Study
These subjects were treated comprehensively by
Barr (1973), and we discuss here only those aspects
needing further emphasis or refinement.
1. Collecting and Extracting Techniques
Thus far, field work on water mites in North
America has focussed largely on qualitative sampling to
obtain specimens required for systematic or life history
studies, and the methods outlined below were developed primarily for these purposes. Efforts to modify
these procedures for rigorous and cost-effective quanti-
573
tative sampling are needed, particularly those aimed at
capitalizing on behavioral traits to simplify extraction
of mites from detritus and silt.
a. Deutonymphs and Adults The best results are
achieved, in most habitats and for virtually all types of
substrates, using a procedure that requires a strong net
with a wide opening (25 – 40 cm diameter) and fine
mesh size (250 m), a supply of strong, clear polyethylene bags (4 – 5 kg capacity), a set of standard sieves of
various mesh size (ranging from 250 m to 2 mm), six
large, leak proof polyethylene containers (1 L), an ice
chest, six rectangular, white photographic trays (30 cm
40 cm), a flashlight, a set of eye droppers with a
range of different sized tip openings, and a supply of
small, leak proof polyethylene containers (50 mL).
Once the desired habitat or microhabitat has been
selected, the substrate must be thoroughly stirred. In
flowing water, the net should be positioned and secured
so that the current will carry dislodged organisms and
particles of substrate into it. In lentic habitats, it must
be swept back and forth through the column of mixed
water and substrate to pick up the organisms.
In very shallow habitats, such as seepage areas,
small rheocrenes, fens, and the edges of pools and
ponds, the substrate must be gathered and stirred by
hand. The net is worked beneath the edge of the wet
moss, leaf litter, or detritus that is to be sampled so that
mats of wet substrate can be separated and rinsed over
the opening. Where there is sufficient surface water, successive handfuls of loose material are picked or scooped
up with the aid of a small trowel, thoroughly picked
apart and washed in the net. Larger pieces of detritus
are discarded after careful washing. In seepage areas,
spring edges, fens, and bog margin habitats where there
is often an extensive carpet of wet moss and associated
plants, mites can also be collected by methodically and
vigorously treading down the plants while dragging the
net along to scoop up the detritus, silt, and organisms
that become temporarily dislodged as water is forced up
through the disturbed layer of vegetation. In either case,
when a mixture of fine silt, plant fragments and organisms begins to fill the bottom of the net, it is transferred
to a plastic bag containing a small amount of water.
This process is repeated until the bag is approximately
half full of material. The bag is then nearly filled with
clean water and the contents stirred gently but thoroughly to allow heavy inorganic material such as gravel
and sand to settle to the bottom. The contents of the
bag are then poured carefully through a set of sieves so
that gravel and sand remain in the bag. This operation
is repeated until all loose organic material is in the
sieves. We find that a pair of upper and lower sieves
with mesh sizes of 1.4 mm and 250 m, respectively,
574
Ian M. Smith, David R. Cook, Bruce P. Smith
yields good results. The coarse material in the upper
sieve is thoroughly stirred and rinsed with clean water.
The fine silt and small organisms accumulating in the
lower sieve are regularly transferred to a large polyethylene container with enough water to just cover the surface of the silt. When the container is about half full of
fine silt it is closed and placed in an ice chest to be
transported back to the laboratory or sorting area. In
situations where there is insufficient surface water to
permit washing and rinsing of samples on site, accumulations of unsorted substrate material must be gathered,
stored in plastic bags and transported on ice to a source
of clean water for subsequent processing.
With few modifications, this procedure can be used
effectively in other types of habitats. Where the water
column is between 25 cm and 1 m in depth and the bottom is firm enough to permit wading, the feet, hands,
and net can be used to agitate the substrate (with the
aid of a spade when necessary). In stream riffles with a
heterogeneous substrate of rocks, gravel, sand, and detritus, vigorous and persistent agitation is usually necessary to dislodge the large number and variety of mites
that cling tenaciously to rocks or burrow in the silt under larger stones. We get good results by securing the
net approximately 1 – 2 m downstream, and then systematically working through the substrate to loosen and
thoroughly rinse all particles so that organisms are carried into the net. Where well-developed interstitial habitats occur in a stream bed, this approach can be extended to yield rich collections of mites including both
surface and hyporheic species. This is accomplished by
simply digging down through the gravel and sand beneath the riffle zone, removing all large stones, to allow
the current to flush out the pockets of interstitial silty
detritus and the organisms that live there. By digging
out an area of substrate approximately 1 m2 to a depth
of 0.5 m, we are often able to recover good numbers of
many hyporheic species. This technique can often be
used to sample the hyporheic zone in habitats where the
standard Karaman – Chappuis method (Chappuis,
1942; Gledhill, 1982) is not practicable.
Quantitative samples of hyporheic arthropods
comprising large and diverse collections of water mites
have recently been obtained in desert streams in Arizona by driving polyvinyl chloride tubing into the substrate to various depths and using a mechanical pump
to evacuate the resulting wells periodically.
In littoral lentic habitats such as depositional pools
in streams, ponds, and sheltered bays in lakes, the feet
can often be used to thoroughly stir up the top few centimeters of sediment and any rooted aquatic plants that
may be present. A net is then repeatedly swept back
and forth through the entire volume of disturbed water
column to ensure that both swimming and crawling
species are captured. As some mites in lentic habitats
are too large to pass readily through 1.4-mm mesh, we
recommend use of an upper sieve with 2.0-mm mesh
when sampling ponds and lakes.
The procedure outlined here can also be used by
scuba divers and a boat operator to collect mites and
insects from the sublittoral and profundal zones of
lakes down to depths of 30 m or more, at a much
higher level of efficiency than is possible with conventional grabs or dredges.
In the sorting area, a number of white photographic
trays are set up on tables and filled to a depth of 2 – 4
cm with clean water. Once the water has settled, the
contents of one polyethylene container are carefully decanted into each tray, so that the silt spreads out in the
center of the tray but does not reach the edges. As the
mites become active and begin to move around in the
trays, they are picked up individually using an eye dropper and transferred to small polyethylene containers.
Many swimming species are strongly positively phototropic and can be recovered efficiently from trays kept
in bright light. However, many crawling mites and a
considerable number of swimming species are moderately positively or negatively phototropic and become
highly active only at low levels of ambient light. The effectiveness and rate of recovering mites from all types of
habitats can be substantially increased by reducing ambient lighting to low levels indoors or waiting until after
sunset outdoors, and then scanning the trays systematically and periodically with a flashlight beam. An experienced collector can then spot the mites as they move
across a contrasting background provided by either the
darkly colored silt or the white bottom of the tray.
Mites remain active and continue to emerge from silt in
the trays for up to 72 h, as deoxygenation of the water
proceeds at room temperature. For this reason, it is suggested that, whenever possible, mites should be extracted from samples in situations that permit periodic
observation of undisturbed trays over a period of at
least 4 – 6 h with controlled lighting. Under these conditions, mites tend to congregate around the edges of the
trays, especially in the corners, where they can be easily
spotted and picked out.
Although water mites frequently are found in samples collected for quantitative analysis in ecological
studies, they are almost always underrepresented in
terms of both numbers and diversity. This is often because collecting devices such as dredges or sieves are
designed to capture organisms that are larger or more
sedentary than mites. Due to their small size and ability
to cling tenaciously to substrates, mites in lotic habitats
can often resist casual efforts to dislodge them or can
simply pass through nets with coarse mesh. By running
or swimming, mites in hyporheic or lentic habitats can
16. Water Mites (Hydrachnida) and Other Arachnids
avoid capture by traps that are designed to sample instantaneously small areas of superficial substrates. Mite
specimens are frequently overlooked when preserved
samples are sorted because extraction techniques concentrate on larger organisms or depend on flotation
methods which we have found ill-suited for separating
water mites efficiently from samples of detritus.
Water mites can be collected from deeper water
with underwater light traps using either battery-powered incandescent bulbs (Hungerford et al., 1955;
Pieczynski, 1962, 1969; Pieczynski and Kajak, 1965)
or chemoluminescent tubes (Barr, 1979) as light
sources. Barr (1979) recommends use of small,
portable light traps for qualitative sampling in lentic
habitats, and provides a useful assessment of their utility for collecting water mites for various purposes, including quantitative analysis.
Recently, a promising technique has been developed for extracting water mites and other aquatic
arthropods quantitatively from samples of substrate by
inducing them to respond actively to a temperature
gradient (Fairchild et al., 1987). This method has
yielded very good results for samples of baled sphagnum from bog ponds, and could probably be adapted
for use with any type of substrate inhabited by mites.
b. Larvae Free-living larvae are often collected
from aquatic habitats along with deutonymphs and
adults, but these specimens must be preserved in order
to be identified and often cannot be reliably associated
with other instars of the same species. Parasitic larvae
can be recovered from insect hosts collected at or near
the parental habitats. Aquatic hosts are usually captured using a dip net, whereas aerial adults can be
trapped as they emerge from water, netted while
swarming (especially over water at dusk), swept from
vegetation, or attracted to lights at night.
2. Rearing
Larvae needed for taxonomic or life history studies
can be reared from females using the techniques described by Prasad and Cook (1972), Barr (1973), and
Smith (1976a). Fertilized females of many species can
be collected in spring and early summer. Females
needed for rearing should be placed individually in
small dishes or vials containing water and a few fragments of moss or wood as a substrate. The containers
should then be covered and placed in a shaded, wellventilated location where the temperature can be maintained near levels experienced in the natural habitat of
the mites. Mites from cold, stenothermic habitats must
be refrigerated. The rearing containers should be
checked periodically for the presence of egg masses. Females should be preserved immediately after they have
575
laid their eggs to ensure that good-quality specimens are
available for confirming species identifications. Larvae
should be either preserved or slide-mounted directly as
they emerge. Species-level treatment of larvae in certain
families can be found in Prasad and Cook (1972),
Wainstein (1980) and various papers by Smith (1976a,
1978, 1979, 1982, 1983a – f, 1984, 1989d, 1992a).
3. Preservation and Preparation for Study
Deutonymphal and adult mites must be preserved
in modified Koenike’s solution (or GAW), consisting of
5 parts glycerin, 4 parts water, and 1 part glacial acetic
acid, by volume (see Barr, 1973), so that they can be
properly cleared, dissected, and slide-mounted for identification and study. Specimens preserved in alcohol or
other hygroscopic agents become so distorted and brittle that subsequent preparation is difficult or impossible. Deutonymphs and adults must be cleared in either
acetic corrosive or 10% KOH, dissected, and mounted
in glycerine jelly, following the techniques described by
Barr (1973) and Cook (1974).
Reared larvae are often slide-mounted directly, but
can be preserved in 70% alcohol for subsequent
mounting without being seriously damaged. Parasitic
larvae should be preserved with the host in 70% alcohol, although larvae removed from hosts that have
been pinned and dried often can be slide-mounted successfully. Before larvae are removed from hosts, the attachment sites should be noted and recorded. Larvae
should be mounted whole in Hoyer’s medium which
also acts as an efficient clearing agent for these small
specimens (see Barr, 1973).
G. Taxonomic Keys to Genera of Water Mites
in North America
The following keys for larvae and adults permit
identification of specimens to family, subfamily and
genus levels. They are intended for use by nonspecialists and use characters that should be obvious on slidemounted specimens to all careful observers. As a result,
the keys are intentionally artificial in that genera do not
necessarily key out with others belonging to the same
family. The larval key does not include all taxa that appear in the adult key, as larvae of many genera remain
unknown (see Table I). Knowledge of larvae in a number of other genera is based on only a few species. Consequently, the key for larvae will be subject to considerable revision and refinement as larvae of more species
are reared and described. We include the genus Stygothrombium (superfamily Styothrombioidea) in the
keys and in Table 1 because these mites are frequently
collected with hydrachnids and closely resemble them
in morphology, behavior and ecology.
576
Ian M. Smith, David R. Cook, Bruce P. Smith
The keys presented here are designed for use with a
minimum of specimen preparation. Larvae should be
whole-mounted on slides. The legs should be spread
out from the body as much as possible by gently pressing down on the coverslip, as the chaetotaxy of leg segments is frequently used as a key character. Adults require clearing and some dissection prior to slide
mounting. The mouth parts (capitulum and appendages) should be removed to permit examination of
the pedipalps in medial and lateral view. Dorsal and
ventral shields, when present, should be separated to
allow clear viewing of certain key characters.
We follow the convention for numbering the setiform structures on the leg segments of larvae that was
established by Prasad and Cook (1972) and modified
slightly by I.M. Smith (1976a) (see Figs. 6 – 8). The setae are numbered starting proximally on the posterolateral surface, proceeding distally along the dorsal half of
the segment to the end, then returning proximally
along the ventral half of the segment. Exceptions are
made in the cases of solenidia (setiform chemoreceptors, see Figs. 31, 32, arrows) which are numbered
first, that is as “s1” on the genua, tibia III, and the
tarsi, and as “s1” and “s2” on tibiae I and II, and
tarsal eupathids (specialized setae, see Fig. 77, arrow)
which are numbered second, that is as “2” on the tarsi.
Where it has been necessary to use leg chaetotaxy in
the key, the number of solenidia present on a segment
is indicated in brackets (e.g. “2s”) immediately following the information on the number of setae, so that
all setiform structures can be accounted for. We usually
have omitted the empodia and claws from the illustrations of the legs of larvae so that the number and positions of distal setae on the tarsi can be seen clearly. The
names of the segments of the appendages are consistently abbreviated as follows:trochanter, Tr; femur, Fe
(basifemur, BFe; telofemur, TFe); genu, Ge; tibia, Ti;
and tarsus, Ta.
1. Taxonomic Key to Genera for Known Larval Water Mites
1a.
Legs with six movable segments, with basifemur (BFe) and telofemur (TFe) separated (Figs. 9, 21, 40, 48) .......................................2
1b.
Legs with five movable segments, with basifemur and telofemur fused (Figs. 6 – 8, 57, 60, 67) ........................................................15
2a(1a).
Gnathosoma with elaborate camerostome enclosing chelicerae, with pedipalps inserted ventrally (Figs. 11, 13, 14) ; dorsal plate present and bearing only two pairs of setae (verticils — ve and vi) near anterior edge (Figs. 11 and 13); coxal plates III located posteriorly on idiosoma with insertions of legs III located at posterolateral edges (posterior to level of excretory pore) and legs III directed
posteriorly (Figs. 12, 15) .........................................................................................................................................Hydrovolzioidea 3
2b.
Gnathosoma lacking elaborate camerostome; dorsal plate absent (Figs. 30, 36), or present and bearing more than two pairs of setae
with verticils (ve and vi) anteriorly and at least internal scapulars (si) in posterior half near edge (Figs. 27, 29, 33); coxal plates III
located near midlength of idiosoma with insertions of legs III located anterolaterally (anterior to level of excretory pore) and legs III
directed laterally (Fig. 28) ..................................................................................................................................................................4
3a(2a).
Dorsal plate relatively small (Fig. 11); venter with rows of numerous small urstigmata borne between coxal plates I and II (Fig. 12,
arrow); coxal plates I and II separated from one another on each side and from opposite members medially (Fig. 12); pedipalps
massive, with tibia (Ti) bearing four long, thick, curved setae (Fig. 10); setae c3 often foliate (Fig. 11)................................................
................................................................................................................................................................Hydrovolziidae Hydrovolzia
3b.
Dorsal plate relatively large (Fig. 13); venter with urstigmata either inconspicuous or absent (Fig. 15); coxal plates I and II fused together to form single anterior plate (Fig. 15); pedipalps moderate in size, with tibia (Ti) bearing two slightly thickened, straight setae (Fig. 14) .....................................................................................................................................Acherontacaridae Acherontacarus
4a(2b).
Dorsal plate absent (Figs. 30, 36), or small, covering less than one-third length of idiosoma and usually with internal scapular setae
(si) at posterior edge (Figs. 27, 29, 33); lateral eyes borne on separate platelets (Figs. 27, 29, 30, 36); leg tarsi with unmodified claws
and empodium (emp) (Fig. 26)...............................................................................................................................Hydryphantoidea 5
4b.
Dorsal plate large, covering well over one-third length of idiosoma and with internal scapular setae (si) near midlength (Figs. 37,
42, 46, 51); lateral eyes borne on single eye plates (Figs. 37, 42, 46); leg tarsi with claws modified or reduced (Figs. 41, 45, 50) ........
........................................................................................................................................................................................Eylaoidea 11
5a(4a).
Dorsal plate roughly quadrangular, bearing four pairs of setae (ve, vi, se, si) (Figs. 27, 33), or three pairs with external scapulars (se)
borne on separate platelets near posterolateral angles of plate ...........................................................................................................6
5b.
Dorsal plate triangular, fragmented, or absent, bearing fewer than three pairs of setae (Figs. 29, 30, 36)...........................................9
6a(5a).
Claw (movable digit) of chelicerae over one-half length of basal segment (Fig. 17); excretory pore setae (ps1 and ps2) and their alveoli absent (Fig. 16) ..................................................................................................................................Thermacaridae Thermacarus
6b.
Claw (movable digit) of chelicerae less than one-third length of basal segment (Fig. 25); excretory pore setae (ps1 and ps2) or at
least their alveoli, present (Figs. 24, 28, 34) ...............................................................................................Hydryphantidae (in part) 7
7a(6b).
Coxal plates II lacking setae (2b absent) (Fig. 20); urstigmata relatively large (Fig. 20, arrow); pedipalp tibia with terminal clawlike
seta deeply bifurcate (Fig. 18, arrow) and tarsus with only one thickened seta; excretory pore plate absent but setae ps1 and ps2
present (Fig. 20)............................................................................................................................................... Wandesiinae Wandesia
16. Water Mites (Hydrachnida) and Other Arachnids
577
7b.
Coxal plates II bearing setae 2b (Fig. 28); urstigmata relatively small (Fig. 28, arrow); pedipalp tibia with terminal clawlike seta
only slightly bifurcate or undivided, and tarsus with two thickened setae (Fig. 19, arrows); excretory pore plate present; setae ps1
and ps2 present or absent, but their alveoli always present (Fig. 28) ..................................................................................................8
8a(7b).
Medial eye present (Fig. 27, arrow) ........................................Thyadinae Panisus, Panisopsis, Thyas, Thyasides, Euthyas, Zschokkea
8b.
Medial eye reduced (Fig. 33) ................................................................................................................Tartarothyadinae Tartarothyas
9a(5b).
Dorsal plate triangular, bearing scapular setae (se and si), with verticil setae (ve and vi) borne on small platelets flanking apex of
plate (Fig. 29); excretory pore setae present, with ps1 borne on plate (Fig. 23) and ps2 borne on small separate platelets; basal segment of chelicerae massive and longitudinally striate (Fig. 25) ....................Hydryphantidae (in part) Hydryphantinae Hydryphantes
9b.
Dorsal plate fragmented or absent, with verticil and scapular setae borne on paired platelets (Figs. 30, 36); excretory pore setae reduced to vestiges or absent, but their alveoli present (Figs. 24, 34); basal segment of chelicerae relatively slender and smooth
(Fig. 27) ...........................................................................................................................................................................................10
10a(9b).
Medial eye present (Fig. 30, arrow); excretory pore plate well sclerotized and nearly triangular (Fig. 24); pedipalp tarsus with all
setae slender; solenidia on leg tarsi slender (Fig. 31, arrow) .............................................Hydryphantidae (in part) Protziinae Protzia
10b.
Medial eye absent (Fig. 36); excretory pore plate poorly sclerotized and oblong (Fig. 34); pedipalp tarsus with two thickened, bladelike setae (Fig. 35, arrows); solenidia on tarsi of legs I and II very thick (Fig. 32, arrow) .......................Hydrodromidae Hydrodroma
11a(4b).
Coxal plates I, II, and III on each side bearing two, two, and two setae, respectively (Figs. 38, 43); tarsi of legs bearing paired claws
and clawlike empodium (emp)(Fig. 41) ............................................................................................................................................12
11b.
Coxal plates I, II, and III on each side bearing two or one, one, and one setae, respectively (Figs. 44, 53); tarsi of legs bearing two
dissimilar clawlike structures terminally (Fig. 45, 50) ..............................................................................................Limnocharidae 13
12a(11a).
Dorsum of idiosoma nearly covered by single, elongate dorsal plate in unengorged larvae, bearing seven pairs of setae (or their alveoli) (Fig. 37); coxal plates with setae all simple (Fig. 38); excretory pore plate elongate (Fig. 39) .................................Eylaidae Eylais
12b.
Dorsum of idiosoma with three separate plates bearing five, one, and one pairs of setae respectively (Fig. 42); coxal plates with setae
1a, 1b, 2b and 3b blunt and conical in shape (Fig. 43); excretory pore plate obcordate (Fig. 43) .........................Piersigiidae Piersigia
13a(11b).
Dorsal plate covering no more than anterior half of idiosomal dorsum, bearing four pairs of setae (verticils and scapulars) (Figs. 46,
51); coxal plates I bearing two pairs of setae, 1a and 1b (Fig. 44); excretory pore plate diamond-shaped (Fig. 47) or nearly oval
(Fig. 49) ...................................................................................................................................................................Limnocharinae 14
13b.
Dorsal plate covering entire idiosomal dorsum in unengorged larvae, bearing seven pairs of setae, including verticils, scapulars and
three pairs of hysterosomal setae (Fig. 52); coxal plates I bearing one pair of setae (Fig. 53); excretory pore plate pyriform (Fig. 53)
.......................................................................................................................................Rhyncholimnocharinae Rhyncholimnochares
14a(13a).
Dorsal plate (Fig. 46) finely punctate; excretory pore plate relatively small, little larger than excretory pore (Fig. 47); leg tarsi bearing two similar clawlike structures with empodium and single ambulacral claw both slender (Fig. 45) ............................Limnochares
14b.
Dorsal plate (Fig. 51) strongly striate; excretory pore plate relatively large, much larger than excretory pore (Fig. 49); leg tarsi bearing two dissimilar clawlike structures, with empodium wider than single ambulacral claw (Fig. 50)...........................Neolimnochares
15a(1b).
Dorsal plate bearing three pairs of setae (external verticils, scapulars) and an unpaired anteromedial seta (probably representing internal verticils) (Fig. 59, arrow); urstigmata (Ur) stalked (Figs. 59, 61); pedipalp tibiae with terminal clawlike seta four-pronged
(Fig. 63, arrow), leg tarsi bearing paired pectinate claws and simple empodium (Fig. 60) ....................................................................
.................................................................................................................Stygothrombidioidea Stygothrombidiidae Stygothrombium
15b.
Dorsal plate bearing at least four pairs of setae (verticils and scapulars), and in some cases one or more additional pairs of hysterosomal setae, but no unpaired setae (Figs. 56, 64, 68); urstigmata sessile (Figs. 58, 65); pedipalp tibiae with terminal clawlike seta
undivided (Fig. 66, arrow), or deeply bisected (Figs. 55, 71, arrows); leg tarsi bearing paired simple claws (Fig. 175) or lacking
claws (Fig. 54), but always bearing a simple empodium ...................................................................................................................16
16a(15b).
Dorsal plate bearing eight pairs of setae, including verticils, scapulars, c3, and three additional pairs of hysterosomal setae (Fig. 56);
leg tarsi lacking paired claws (Fig. 54); excretory pore plate tiny, and bearing neither setae nor their alveoli (Fig. 58, arrow)..............
...........................................................................................................................................Hydrachnoidea Hydrachnidae Hydrachna
16b.
Dorsal plate bearing four-to-six pairs of setae, including only the verticils and scapulars (Fig. 64) or, in some cases, the verticils,
scapulars and one or two pairs of hysterosomal setae (Fig. 68); leg tarsi bearing paired claws (Fig. 143, 175); excretory pore plate
small to large, always bearing setae ps1 and ps2 or at least their alveoli (Figs. 62, 80, 104, 109, 113 – 114, 172, 176, 185, 193) ....17
17a(16b).
Coxal plates I, II, and III on each side all separate (Figs. 65, 69, 73, 172, 179, 180, 186), or all fused (Fig. 185) and plates of two
sides fused medially and dorsal plate round and bearing setae c1 laterally (Fig. 184) .......................................................................18
17b.
Coxal plates I and II on each side separate and plates II and III fused at least medially (Figs. 83, 106, 145, 157), or all plates on
each side fused (Figs. 92, 100) but plates of two sides separate medially and dorsal plate elliptical and not bearing setae c1 (Figs. 91,
101, 144, 150, 153, 168) .................................................................................................................................................................34
578
Ian M. Smith, David R. Cook, Bruce P. Smith
18a(17a).
Tarsi of legs III bearing 12 or more setae, including Ta8 (Fig. 8, arrow); dorsal plate usually elongate and elliptical and bearing only
four pairs of setae (verticils and scapulars) (Figs. 3, 64, 74); when dorsal plate round and bearing five pairs of setae, including c1
laterally (Fig. 68), then coxal plates III bearing setae pa and h2 in addition to 3a (Fig. 69) and all setae on pedipalp tarsi short and
bladelike (Fig. 71) ..........................................................................................................................................Lebertioidea (in part) 19
18b.
Tarsi of legs III usually bearing 11 or fewer setae, lacking at least Ta8 (Figs. 174, 181, 196, 198); dorsal plate usually nearly round
(Figs. 178, 184, 188, 190, 191), rarely elongate and elliptical (Fig. 171); when tarsi of legs III bearing 12 setae then dorsal plate
round and bearing five pairs of setae, including c1 laterally (Fig. 195) and coxal plates III bearing only setae 3a (Fig. 193) and at
least one seta on pedipalp tarsi long and whip-like (similar to Fig. 177) .....................................................................Arrenuroidea 23
19a(18a).
Pedipalp tarsi with no setae as long as pedipalp (Fig. 71); tarsi of legs I, II, and III bearing 13 or 14 (1s), 13 or 14 (1s), and 12
setae respectively, lacking Ta15 ...................................................................................................................Anisitsiellidae (in part) 20
19b.
Pedipalp tarsi with at least one seta as long as pedipalp (Figs. 2, 66); tarsi of legs I, II, and III bearing 15 (1s), 15 (1s), and 13 or
14 setae respectively, including Ta15 (Figs. 6 – 8, 67)........................................................................................................................21
20a(19a).
Dorsal plate bearing six pairs of setae, including verticils, scapulars and both c1 and e1 on lateral edge, and with setae si long,
thick, and located near midlength (Fig. 68); coxal plates III bearing three pairs of setae, including 3a and both pa and h2 near posterior edges (Fig. 69) ..........................................................................................................................................................Bandakiopsis
20b.
Dorsal plate bearing four pairs of setae (verticils and scapulars), with setae si short, slender, and located in anterior third of plate
(Fig. 70); coxal plates III bearing only setae 3a (Fig. 72) ........................................................................................................Utaxatax
21a(19b).
Dorsal plate longitudinally striate (Figs. 3, 64); coxal plates III bearing only setae 3a (Figs. 4, 65); excretory pore plate small and
usually quadrangular (Figs. 5, 65) ............................................................................................................................Sperchontidae 22
21b.
Dorsal plate reticulate (Fig. 74); coxal plates III bearing two pairs of setae, 3a and supernumerary setae 3b (Fig. 73); excretory pore
plate relatively large and diamond-shaped (Fig. 73) ............................................................................................Teutoniidae Teutonia
22a(21a).
Coxal plates II bearing one pair of setae, 2b, posterolaterally (Fig. 65) .................................................................................Sperchon
22b.
Coxal plates II lacking setae (Fig. 4)...............................................................................................................................Sperchonopsis
23a(18b).
Coxal plates I, II, and III on each side all fused and coxal plates of two sides fused medially, with setae 1a, 2b and 3a reduced to
vestiges (Fig. 185); pedipalp tarsi with no setae as long as pedipalp (Fig. 182); dorsal plate with setae si reduced to vestiges (Fig.
184) ....................................................................................................................................................Acalyptonotidae Acalyptonotus
23b.
Coxal plates I, II, and III on each side all separate and coxal plates of two sides separate (Figs. 172, 186, 189, 192, 193); pedipalp
tarsi with at least one seta as long as pedipalp (Fig. 177) .................................................................................................................24
24a(23b).
Idiosoma moderately flattened dorsoventrally; gnathosoma projecting beyond anterior edge of dorsal plate, entirely exposed in dorsal view (Figs. 171, 195); excretory pore plate variously shaped (Figs. 172, 193, 197) but usually not attenuate anteriorly; pedipalp
tarsi with long, thick, distal seta straight or only slightly bowed basally; leg genua with setae Ge5 borne on tubercles that usually
are prominent (Fig. 173, arrow) .......................................................................................................................................................25
24b.
Idiosoma extremely flattened dorsoventrally; gnathosoma recessed beneath anterior edge of dorsal plate, partially or entirely concealed in dorsal view (in unmounted specimens) (Fig. 188); excretory pore plate triangular with anterior apex attenuate (Figs. 176,
192); pedipalp tarsi with long, thick, distal seta strongly bowed, and usually lobed, basally (Figs. 177, 183, arrows); leg genua with
setae Ge5 not borne on tubercles......................................................................................................................................................29
25a(24a).
Dorsal plate bearing four pairs of setae (verticils and scapulars), with setae c1 on lateral membranous integument (Fig. 171); setae
c2 and d2 thick and long relative to other hysterosomal setae (Fig. 171); excretory pore setae ps1 and ps2 absent, represented by
their alveoli on excretory pore plate (Fig. 172); tibiae of legs I bearing eight setae (2s), including Ti11; leg tarsi lacking setae Ta14
(Figs. 173, 175) ...........................................................................................................................................................Momoniidae 26
25b.
Dorsal plate bearing five pairs of setae, including the verticils, scapulars and setae c1 laterally (Fig. 195); setae c2 and d2 similar to
other hysterosomal setae (Fig. 195); excretory pore setae ps1 and ps2 present on plate (Fig. 193, 197), tibiae of legs I bearing seven
setae (2s), lacking Ti11; leg tarsi bearing setae Ta14 (Figs. 196, 198, arrows) ...............................................................................27
26a(25a).
Legs with setae Tr1, Ge5, Ti9 and Ti11 much longer than respective segments; setae IITi9, IITi11, IIIGe5, IIITi9 and IIITi11
plumose (Fig. 173) ...........................................................................................................................................Momoniinae Momonia
26b.
Legs with setae Tr1, Ge5, Ti9 and Ti11 shorter than respective segments; setae IITi9, IITi11, IIIGe5, IIITi9 and IIITi11 bladelike
(Fig. 175) ........................................................................................................................................Stygomomoniinae Stygomomonia
27a(25b).
Tarsi of legs III bearing nine setae, lacking Ta12 (Fig. 198); tarsi of legs II and III with setae Ta4 and Ta6 long (usually as long as
respective segments) (Figs. 198); coxal plates I and II with conspicuous denticulate projections on posterior edges (Figs. 197, 199,
arrows), coxal plates III with transverse muscle attachment scars (TMAS), excretory pore plate subtriangular (Figs. 197, 199) .........
..............................................................................................................................................................................Krendowskiidae 28
27b.
Tarsi of legs III bearing at least 11 setae, including Ta12 (Fig. 196); tarsi of legs II and III with setae Ta4 and Ta6 usually short (conspicuously shorter than respective segments) (Fig. 196); coxal plates I and II lacking denticulate projections on posterior edges,
16. Water Mites (Hydrachnida) and Other Arachnids
579
coxal plates III lacking transverse muscle attachment scars (TMAS), excretory pore plate variously shaped, but rarely triangular
(Fig. 193) ..........................................................................................................................................................Arrenuridae Arrenurus
28a(27a).
Coxal plates I and II each with row of denticulate projections along whole posterior edge (Fig. 197, arrows); TMAS weakly developed and short (Fig. 197); tubercles bearing setae f2 large, prominent, and medially attenuate (Fig. 197) ......................Krendowskia
28b.
Coxal plates I and II each bearing only few, denticulate projections in medial third of posterior edge (Fig. 199); TMAS strongly developed and long (Fig. 199); tubercles bearing setae f2 not unusually prominent (Fig. 199) ......................................................Geayia
29a(24b).
Dorsal plate bearing five pairs of setae, including verticils, scapulars and setae c1 laterally (Fig. 188) .............Mideopsidae Mideopsis
29b.
Dorsal plate bearing four pairs of setae (verticils and scapulars), with setae c1 on lateral membranous integument (Figs. 178, 190)
.........................................................................................................................................................................................................30
30a(29b).
Tibiae of legs II and III bearing nine setae (2s and 1s, respectively), including both Ti10 and Ti11 (Figs. 174, 181) ...................31
30b.
Tibiae of legs II and III bearing fewer than nine setae (2s and 1s, respectively), lacking either Ti10 or Ti11, or both (Fig. 187,
194) .................................................................................................................................................................................................32
31a(30a).
Pedipalp tarsi with long, thick, distal seta bowed, but not deeply lobed or fringed basally, and with most medial seta moderately
thick, but not fringed (Fig. 183); legs II and III with setae Ge5, Ti9 and Ti11 much longer than respective segments and plumose,
and setae Ta4 and Ta6 much longer than respective segments (Fig. 181).....................................................................Mideidae Midea
31b.
Pedipalp tarsi with long, thick, distal seta bowed, deeply lobed and fringed basally, and most medial seta thick and fringed (Fig.
177); legs II and III with setae Ge5, Ti9 and Ti11 shorter than respective segments and simple, and setae Ta4 and Ta6 shorter than
respective segments (Fig. 174) ...............................................................................Nudomideopsidae Nudomideopsis, Paramideopsis
32a(30b).
Tibiae of legs II bearing seven setae (2s), lacking Ti10 and Ti11 (Fig. 194); tibiae of legs III bearing eight setae (1s), lacking Ti10
.............................................................................................................................................................................Laversiidae Laversia
32b.
Tibiae of legs II and III bearing eight setae (2s and 1s respectively), including Ti10, but lacking Ti11 (Fig. 187) ........................33
33a(32b).
Dorsum with setae si located well in anterior half of plate and setae c1 located in lateral integument near midlength of plate, posterior to level of setae si (Fig. 191) ..............................................................................................Neoacaridae Neoacarus, Volsellacarus
33b.
Dorsum with setae si near midlength of plate and setae c1 located in lateral integument near setae c3, anterior to level of setae si
(Fig. 190) ...............................................................................Athienemanniidae Athienemanniinae Chelomideopsis, Platyhydracarus
34a(17b).
Coxal plates III bearing setae 3a and at least one other pair of setae (pa or 3b) and excretory pore plate bearing only setae ps1 and
ps2 (Figs. 83, 86, 90, 92) ..............................................................................................................................Lebertioidea (in part) 35
34b.
Coxal plates III usually bearing only one pair of setae, 3a (Figs. 80, 97, 100, 145, 152), when coxal plates III also bearing setae pa,
then excretory pore plate bearing setae h2 in addition to ps1 and ps2 (Figs. 104, 159 – 163, 166) ...................................................38
35a(34a).
Coxal plates III truncate posteriorly, bearing three pairs of setae, including 3a and both pa and h2 on posterior edges (Fig. 83); tibiae of legs I, II, and III bearing nine setae (2s, 2s, and 1s, respectively), including both Ti9 and Ti10 (Fig. 82); tarsi of legs I
and II bearing 14 setae (1s), including Ta14 and Ta15 (Fig. 82); cheliceral bases separate (Fig. 81, arrow) .......................................
...........................................................................................................................................................Lebertiidae Lebertia, Estelloxus
35b.
Coxal plates III rounded or pointed posteriorly, bearing two pairs of setae, including setae 3a and either 3b laterally or pa posteromedially (Figs. 86, 90, 92); tibiae of legs I, II, and III bearing eight setae (2s, 2s, and 1s, respectively), lacking Ti9 or Ti10
(Figs. 85, 88, 89); tarsi of legs I and II bearing 13 setae (1s), lacking Ta15 (Fig. 88), or 12 setae (1s), lacking both Ta15 and
Ta14 (Fig. 85); cheliceral bases fused (Figs. 84, 87, 91) ....................................................................................................................36
36a(35b).
Dorsal plate bearing five pairs of setae including verticils, scapulars and setae c1 laterally; integument beneath posterior edge of dorsal plate intricately folded (Fig. 84, arrow); coxal plates I elongate, extending posteriorly nearly to level of excretory pore plate;
coxal plates III bearing setae 3b laterally (Fig. 86); tarsi of legs I and II bearing 12 setae (1s), lacking Ta14 (Fig. 85) .......................
....................................................................................................................................................................Oxidae Oxus, Frontipoda
36b.
Dorsal plate bearing four pairs of setae (verticils and scapulars), or five pairs including verticils, scapulars and setae d1 laterally; integument beneath posterior edge of dorsal plate not intricately folded (Figs. 87, 91); coxal plates I not elongate, extending posteriorly only to level of insertion of legs III (Fig. 90), or fused with plates II (Figs. 92); plates III bearing setae pa posteromedially (Figs.
90, 92); tarsi of legs I and II bearing 13 setae (1s), including Ta14 (Fig. 88) ........................................................Torrenticolidae 37
37a(36b).
Dorsal plate bearing five pairs of setae including verticils, scapulars and setae d1 laterally, and with setae se long and thick (Fig. 87);
coxal plates I clearly delineated by complete suture lines, and plates II and III fused with suture line incomplete medially (Fig. 90);
excretory pore plate with convex projection posteromedially (Fig. 90); tibiae of all legs lacking setae Ti10, genua of legs III bearing
four setae (1s) including Ge3, and tarsi of legs III bearing 11 setae, lacking Ta10 (Fig. 89)...................Testudacarinae Testudacarus
37b.
Dorsal plate bearing only four pairs of setae (verticils and scapulars), and with setae se relatively short and slender (Fig. 91); coxal
plates of each side all fused, with suture lines obliterated except laterally (Fig. 92); excretory pore plate without projection posteromedially (Fig. 92); tibiae of all legs lacking setae Ti9, genua of legs III bearing three setae (1s), lacking Ge3, tarsi of legs III bearing
12 setae, including Ta10 (Fig. 93).............................................................................................................Torrenticolinae Torrenticola
580
Ian M. Smith, David R. Cook, Bruce P. Smith
38a(34b).
Cheliceral bases separate (Fig. 78, arrow); tarsi of legs III bearing 12 setae, including Ta9 (Fig. 79, arrow); excretory pore plate very
large (equal in width to coxal plates of one side), bearing setae ps1 and ps2 anteromedially and occasionally also setae h2 at posterolateral angles (Figs. 75, 80), and coxal plates I separate from posterior coxal groups, and coxal plates III bearing only setae 3a
(Fig. 80)......................................................................Lebertioidea (in part) Anisitsiellidae (in part) Anisitsiellinae (in part) Bandakia
38b.
Cheliceral bases fused (Figs. 101, 107, 108, 144, 150); tarsi of legs III usually bearing 11 or fewer setae, lacking Ta9 (Fig. 132) (exception Limnesiidae); excretory pore plate small to very large, when equal in width to coxal plates of one side (measured from midline to insertion of leg III) (Figs. 100, 103, 104, 152, 164) then either excretory pore located near posterior edge of plate (Fig. 152,
arrow), or coxal plates I fused to posterior coxal groups (Fig. 100), or coxal plates III bearing setae pa in addition to setae 3a (Figs.
164) .........................................................................................................................................................................Hygrobatoidea 39
39a(38b).
Two pairs of urstigmata borne distally between coxal plates I and II (Figs. 97, arrows A; Fig. 98, arrows); tibiae of legs I and II
bearing seven or eight setae (2s), including only three ventral setae, lacking either Ti8 and Ti9 or Ti10 and Ti11 (Figs. 95, 99) .......
.....................................................................................................................................................................................Limnesiidae 40
39b.
One pair of urstigmata borne distally between coxal plates I and II (Figs. 100, arrow, 106, 111, 133, 164); tibiae of legs I and II
usually bearing nine or more setae (2s), including five ventral setae (Ti7, Ti8, Ti9, Ti10, and Ti11) (Figs. 102, 141, 149) (exception Feltriidae)..................................................................................................................................................................................41
40a(39a).
Coxal plates I and II completely separate (Fig. 98); tarsi of legs III bearing nine setae, lacking Ta9, Ta13 and Ta14 (Fig. 99); dorsal
plate extending laterally well beyond level of eye plates (contrast with Fig. 94)....................................................Tyrrelliinae Tyrrellia
40b.
Coxal plates I and II on each side fused, with suture lines obliterated medially (Fig. 97, arrow B); tarsi of legs III bearing 12 setae,
including Ta9, Ta13 and Ta14 (Fig. 96); dorsal plate narrow, usually confined to region between eye plates (Fig. 94) .........................
...........................................................................................................................................................................Limnesiinae Limnesia
41a(39b).
Coxal plates I, II, and III on each side all fused (Fig. 100)..........................................................................................Hygrobatidae 42
41b.
Coxal plates I separate from posterior coxal group on each side (Figs. 106, 111, 145, 152, 169) ....................................................43
42a(41a).
Excretory pore plate bearing three pairs of setae, including ps1 and ps2 medially and h2 near anterolateral angles, with setae pa on
posterior edges of coxal plates III (Fig. 104) .......................................................................................................................Hygrobates
42b.
Excretory pore plate bearing four pairs of setae, including ps1 and ps2 medially, pa near anterior edge, and h2 near lateral angles
(Figs. 100, 103) ....................................................................................................................................................................Atractides
43a(41b).
Tarsi of legs I and II bearing 10 setae (1s), lacking Ta14, and either Ta12 (Fig. 105) or Ta9 (Figs. 112, 116) ................................44
43b.
Tarsi of legs I and II bearing 11 or more setae (1s), including Ta9 (Figs. 128, 170, arrows; Figs. 147, 151, arrows A) ..................47
44a(43a).
Dorsal and coxal plates, and leg sclerites, conspicuously longitudinally striate (as in Fig. 107); coxal plates III rounded posteriorly
(Fig. 106); tibiae of legs I bearing seven setae (2s), lacking Ti10 and Ti11 (Fig. 105) ..............................................Feltriidae Feltria
44b.
Dorsal and coxal plates reticulate (Fig. 108); coxal plates III bearing pointed projections posteriorly (Fig. 111, arrow); tibiae of legs
I bearing 9 setae (2s), including Ti10 and Ti11 (Fig. 112, 116) ..............................................................................Unionicolidae 45
45a(44b).
Tarsi of legs with empodium undivided (Fig. 115) .......................................................................................................Pionatacinae 46
45b.
Tarsi of legs with empodium bifurcate (Fig. 110, arrow) .............................................................................Unionicolinae Unionicola
46a(45a).
Excretory pore plate usually oval in shape with excretory pore borne near anterior edge (Fig. 114) ....................................Neumania
46b.
Excretory pore plate obcordate with excretory pore borne near middle of plate (Fig. 113)....................................................Koenikea
47a(43b).
Tibiae of all legs bearing seven setae (2s, 2s, and 1s respectively), lacking Ti9 and Ti11 (Fig. 170) ..................Aturidae (in part)
...................................................................................................................................................................................Aturinae Aturus
47b.
Tibiae of all legs bearing eight or more setae (2s, 2s, and 1s, respectively), including Ti9 and Ti11 (Fig. 151).........................48
48a(47b).
Coxal plates III with pointed or lobed projections posteriorly, bearing two pairs of setae, 3a anteromedially and pa posteromedially
(Figs. 156, 164); excretory pore plate large, bearing three pairs of setae, ps1 and ps2, and h2 posterolaterally (Figs. 159 – 163, 166)
.........................................................................................................................................Aturidae (in part) Axonopsinae (in part) 49
48b.
Coxal plates III without, or with small, projections posteriorly, bearing only setae 3a (Figs. 117, 145, 152, 154); excretory pore
plate small or large, bearing only setae ps1 and ps2 (Figs. 118, 123, 135, 138, 142, 146)................................................................52
49a(48a).
Tarsi of legs I and II bearing 12 setae, including setae Ta8 (Fig. 158); excretory pore plate with setae ps1 at or near anterior edge
(Figs. 159, 160) ................................................................................................................................................................................50
49b.
Tarsi of legs I and II bearing 11 setae, lacking setae Ta8 (Fig. 167); excretory pore plate with setae ps1 near anterior edge to posterior to midlength (Figs. 162, 163, 166) ............................................................................................................................................51
50a(49a).
Excretory pore plate subtriangular (Fig. 159) with acutely rounded posterolateral angles, with setae ps2 anterior to midlength between setae ps1 and h2; dorsal plate not reticulate; coxal plates I weakly fused medially with plates II, and with setae 1a long, extending posteriorly beyond level of setae 3a (Fig. 157) ......................................................................................................Estellacarus
16. Water Mites (Hydrachnida) and Other Arachnids
581
50b.
Excretory pore plate obcordate (Fig. 160) or nearly quadrangular (Fig. 161), with broadly rounded posterolateral angles, and with
setae ps2 posterior to midlength, not between setae ps1 and h2; dorsal plate reticulate; coxal plates I separate from plates II, and
with setae 1a relatively short, not extending posteriorly beyond level of setae 3a (Fig. 156) ...........................................Woolastookia
51a(49b).
Suture line between coxal plates II and III long, extending medially to near base of setae 3a (Fig. 164); excretory pore plate either
moderately large and subtriangular (Fig. 162) or very large and subquadrangular (Fig. 163) ............................................Brachypoda
51b.
Suture line between coxal plates II and III short, not extending medially to region of base of setae 3a (Fig. 165); excretory pore plate
moderately large and broadly elliptical or subquadrangular (Fig. 166) ................................................................................Axonopsis
52a(48b).
Coxal plates III with small projections posteriorly (Fig. 117, arrow); excretory pore plate small, little larger than combined area of
excretory pore and bases of setae ps1 and ps2 (Fig. 118)........................................................................................Wettinidae Wettina
52b.
Coxal plates III without projections posteriorly (Figs. 124, 133, 137, 140, 145, 152, 154); excretory pore plate considerably larger
than combined area of excretory pore and bases of setae ps1 and ps2 (Figs. 123, 125, 129, 138, 142, 152) ....................................53
53a(52b).
Tarsi of legs I bearing 13 setae (1s), including Ta8 (Figs. 147, 151, arrows B); excretory pore plate large, and usually triangular or
obcordate (Figs. 138, 139, 148, 152) ...............................................................................................................................................54
53b.
Tarsi of legs I bearing 12 setae (1s), lacking Ta8 (Fig. 128); excretory pore plate small to large, and variously shaped (Figs. 119,
121, 123, 125, 134, 135) .................................................................................................................................................................58
54a(53a).
Excretory pore plate with setae ps1 close together adjacent to excretory pore in posterior half of plate (Figs. 138, 139, 142, 146,
148) ....................................................................................................................................................................Pionidae (in part) 55
54b.
Excretory pore plate with setae ps1 widely separated and removed from excretory pore in anterior half of plate (Fig. 154, arrow)
...........................................................................................................................................................................Aturidae (in part) 57
55a(54a).
Tarsi of legs II bearing 13 setae (1s), including Ta8 (Fig. 141); tibiae of legs with setae Ti9 and Ti11 simple (Fig. 141) ....................
.....................................................................................................................................................................Najadicolinae Najadicola
55b.
Tarsi of legs II usually bearing 12 setae (1s), lacking Ta8 (Fig. 149), when bearing 13 setae then tibiae of legs with Ti9 and Ti11
long and plumose (as in Fig. 149) .....................................................................................................................................Pioninae 56
56a(55b).
Femora of legs II with Fe5 shorter than segment; excretory pore plate usually much less than twice as wide as long, when nearly
twice as wide as long then plate is concave or transverse anteromedially (Fig. 146) ....................................................................Piona
56b.
Femora of legs II with Fe5 longer than segment; excretory pore plate twice as wide as long and convex anteromedially (Fig. 139)
........................................................................................................................................................................................Nautarachna
57a(54b).
Excretory pore plate broadly obcordate (Fig. 152) ..................................................................................Axonopsinae (in part) Ljania
57b.
Excretory pore plate triangular (Fig. 154) ......................................................................................................................Albiinae Albia
58a(53b).
Excretory pore plate small and subtriangular, with setae ps2 borne near posterolateral angles of plate (Fig. 155); suture lines between coxal plates II and III parallel to anterior edge of plate II, coxal plates without distinct lateral coxal apodemes and coxal
plates III lacking medial coxal apodemes and transverse muscle attachment scars (Fig. 155) ....................................Aturidae (in part)
................................................................................................................................................Axonopsinae (in part) Neobrachypoda
58b.
Excretory pore plate variously shaped, when subtriangular with setae ps2 borne near posterolateral angles of plate then suture lines
between coxal plates II and III terminating in distinct lateral coxal apodemes (LCA) that are usually nearly transverse and coxal
plates III bearing at least medial coxal apodemes (MCA) (Figs. 120, 124, 126, 129, 133, 136)...........................Pionidae (in part) 59
59a(58b).
Suture line between coxal plates II and III on each side extending medially beyond lateral coxal apodeme nearly to midline
(Fig. 120, arrow); excretory pore borne on elevated projection which extends to or beyond posterior edge of plate (Figs. 119, 121)
......................................................................................................................................................Hydrochoreutinae Hydrochoreutes
59b.
Suture line between coxal plates II and III on each side extending medially only as far as lateral coxal apodeme (Figs. 122, 124, 126,
129, 133, 136); excretory pore usually not borne on elevated projection, when on small projection this does not extend to posterior
edge of plate .....................................................................................................................................................................................60
60a(59b).
Trochanters of legs III bearing two setae, including supernumerary dorsal seta (Fig. 131, arrow), or tibiae of legs II and III bearing 11 or 12 setae (2s and 1s, respectively), including two supernumerary posteroventral setae (Fig. 132, arrows), or coxal
plates III lacking transverse muscle attachment scars and excretory pore plate with setae ps2 near posterior edge of plate (Fig. 136)
.......................................................................................................................................................................................Foreliinae 61
60b.
Trochanters of legs III bearing only one seta; tibiae of legs II and III bearing only nine setae (2s and 1s, respectively) (as in Figs.
141, 149); coxal plates III usually with transverse muscle attachment scars (Figs. 124, 126, 133), when coxal plates lacking these
scars (as in Fig. 122) then excretory pore plate with setae ps2 removed from posterior edge of plate (Fig. 123)...............................62
61a(60a).
Trochanters of legs III bearing two setae, including supernumerary dorsal seta (Fig. 131, arrow) ....................................Pseudofeltria
61b.
Trochanters of legs III bearing one seta......................................................................................................................................Forelia
582
Ian M. Smith, David R. Cook, Bruce P. Smith
62a(60b).
Excretory pore plate transversely elliptical or semicircular (Fig. 123) ...........................................................Huitfeldtiinae Huitfeldtia
62b.
Excretory pore plate broadly elliptical, subcircular, or subtriangular ...............................................................................Tiphyinae 63
63a(62b).
Excretory pore plate three times as wide as long and broadly elliptical or dumbbell shaped (Figs. 124, 125) .......................Neotiphys
63b.
Excretory pore plate less than twice as wide as long and subcircular (Figs. 127, 130) to subtriangular.....................Pionopsis, Tiphys
FIGURES 2–5
Sperchonopsis ecphyma Prasad and Cook (Sperchontidae), larva. Fig. 2, Venter of
gnathosoma; Fig. 3, dorsum of idiosoma and gnathosoma; Fig. 4, venter of idiosoma; Fig. 5, excretory pore
plate. See text for nomenclature and abbreviations of setae.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 6–8 Sperchonopsis ecphyma Prasad and Cook (Sperchontidae), larva, anterolateral views of legs.
Fig. 6, Leg I; Fig 7, Leg. II; and Fig. 8, Leg III. Fe, Femur; Ge, genu; Ta, tarsus; Ti, tibia; Tr, trochanter.
583
584
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 9–12
Hydrovolzia gerhardi Mitchell (Hydrovolziidae), larva. Fig. 9, Leg I, anterolateral view;
Fig. 10, distal segments of pedipalp; Fig. 11, dorsum of idiosoma and gnathosoma; Fig 12, venter of idiosoma.
FIGURES 13–15 Acherontacarus sp. (Acherontacaridae), larva. Fig. 13, Dorsum of idiosoma and gnathosoma; Fig. 14, venter of gnathosoma; Fig. 15, venter of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 16–17
Thermacarus nevadensis Marshall (Thermacaridae), larva. Fig. 16, Excretory pore plate;
Fig 17, chelicera. FIGURE 18, 20 Wandesia sp. (Wandesiinae), larva. Fig. 18, Tibia and tarsus of pedipalp;
Fig. 20, venter of idiosoma and gnathosoma. FIGURES 19, 21 – 22, 26 – 28 Thyas stolli Koenike
(Thyadinae), larva. Fig. 19, Tarsus of pedipalp; Fig. 21, leg I; Fig. 22, pedipalp; Fig. 26, claws and empodium;
Fig. 27, dorsum of idiosoma and gnathosoma; Fig. 28, venter of idiosoma and gnathosoma. FIGURES 23, 25,
29 Hydryphantes ruber (de Geer) (Hydryphantinae), larva. Fig. 23, Excretory pore plate; Fig. 25, chelicerae;
Fig. 29, dorsum of idiosoma and gnathosoma. FIGURE 24 Protzia sp. (Protziinae), larva, excretory
pore plate.
585
586
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 30 – 31 Protzia sp. (Protziinae), larva. Fig. 30, Prodorsal region of idiosoma; Fig. 31, tibia and
tarsus of leg I. FIGURES 32, 34 – 36 Hydrodroma despiciens (Müller) (Hydrodromidae), larva. Fig. 32,
Tarsus of leg I; Fig. 34, excretory pore plate; Fig. 35, tarsus of pedipalp; Fig. 36, dorsum of idiosoma and
gnathosoma. FIGURE 33 Tartarothyas sp. (Tartarothyadinae), larva, dorsal plate. FIGURES 37 – 41 Eylais
major Lanciani (Eylaidae), larva. Fig. 37, Dorsum of idiosoma and gnathosoma; Fig. 38, venter of idiosoma
and gnathosoma; Fig. 39, excretory pore plate; Fig. 40, leg I; Fig. 41, claws and empodium of leg I.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 42 – 43 Piersigia sp. (Piersigiidae), larva. Fig. 42, Dorsum of idiosoma; Fig. 43, venter of idiosoma
and gnathosoma. FIGURES 44 – 48 Limnochares americana Lundblad (Limnocharinae), larva. Fig. 44, venter
of idiosoma and gnathosoma; Fig. 45, claw and empodium of leg I; Fig. 46, dorsum of idiosoma and gnathosoma; Fig. 47, excretory pore plate; Fig. 48, leg I. FIGURES 49 – 51 Neolimnochares sp. (Limnocharinae),
larva. Fig. 49, Excretory pore plate; Fig. 50, claw and empodium of leg I; Fig. 51, dorsal plate.
587
588
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 52 – 53
Rhyncholimnochares kittatinniana Habeeb (Rhyncholimnocharinae), larva. Fig. 52,
Dorsum of idiosoma and gnathosoma; Fig. 53, venter of idiosoma. FIGURES 54 – 58 Hydrachna
magniscutata Marshall (Hydrachnidae), larva. Fig. 54, Empodium of leg I; Fig. 55, distal segment of pedipalp;
Fig. 56, dorsum of idiosoma and gnathosoma; Fig. 57, leg I; Fig. 58, venter of idiosoma and gnathosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 59 – 61, 63 Stygothrombium sp. (Stygothrombidiidae), larva. Fig. 59, Dorsum of idiosoma and
gnathosoma; Fig. 60, leg I; Fig. 61, venter of idiosoma; Fig. 63, tibia and tarsus of pedipalp. FIGURES 62,
64 – 67 Sperchon glandulosus Koenike (Sperchontidae), larva. Fig. 62, Excretory pore plate; Fig. 64, dorsum
of idiosoma and gnathosoma; Fig. 65, venter of idiosoma; Fig. 66, pedipalp; Fig. 67, leg I.
589
590
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 68 – 69, 71
Bandakiopsis fonticola Smith (Anisitsiellidae), larva. Fig. 68, Dorsum of idiosoma and
gnathosoma; Fig. 69, venter of idiosoma; Fig. 71, pedipalp. FIGURES 70,72 Utaxatax newelli Habeeb
(Anisitsiellidae), larva. Fig. 70, dorsum of idiosoma and gnathosoma; Fig. 72, venter of idiosoma. FIGURES
73, 74 Teutonia lunata Marshall (Teutoniidae), larva. Fig. 73, Venter of idiosoma; Fig. 74, dorsum of
idiosoma and gnathosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 75 – 80 Bandakia phreatica Cook (Anisitsiellidae), larva. Fig. 75, Excretory pore plate; Fig. 76,
pedipalp; Fig. 77, tibia and tarsus of leg I; Fig. 78, dorsum of idiosoma and gnathosoma; Fig. 79, tibia and
tarsus of leg III. Fig. 80, venter of idiosoma. FIGURES 81 – 83 Lebertia sp. (Lebertiidae), larva. Fig. 81,
Dorsum of idiosoma and gnathosoma; Fig. 82, tibia and tarsus of leg I; Fig. 83, venter of idiosoma.
591
592
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 84 – 86 Oxus elongatus Marshall (Oxidae), larva. Fig. 84, Dorsum of idiosoma and gnathosoma;
Fig. 85, tibia and tarsus of leg I; Fig. 86, venter of idiosoma. FIGURES 87 – 90 Testudacarus sp.
(Testudacarinae), larva. Fig. 87, Dorsum of idiosoma and gnathosoma; Fig. 88, tibia and tarsus of leg I;
Fig. 89, distal segments of leg III; Fig. 90, venter of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 91 – 93 Torrenticola sp. (Torrenticolidae), larva. Fig. 91, dorsum of idiosoma and gnathosoma;
Fig. 92, venter of idiosoma; Fig. 93, distal segments of leg III. FIGURES 94 – 97 Limnesia marshalliana
Lundblad (Limnesiinae), larva. Fig. 94, dorsum of idiosoma and gnathosoma; Fig. 95, tibia and tarsus of leg II;
Fig. 96, tibia and tarsus of leg III; Fig. 97, venter of idiosoma and gnathosoma.
593
594
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 98 – 99 Tyrrellia sp. (Tyrrelliinae), larva. Fig. 98, Venter of idiosoma; Fig. 99, tibia and tarsus of
leg III. FIGURES 100 – 101, 103 Atractides grouti Habeeb (Hygrobatidae), larva. Fig. 100, Venter of
idiosoma and gnathosoma; Fig. 101, dorsum of idiosoma and gnathosoma; Fig. 103, excretory pore plate.
FIGURE 102 Hygrobates sp. (Hygrobatidae), larva, tibia and tarsus of leg II. FIGURE 104 Hygrobates
neocalliger Habeeb (Hygrobatidae), larva, excretory pore plate region. FIGURES 105 – 107 Feltria sp.
(Feltriidae), larva. Fig. 105, Tibia and tarsus of leg I; Fig. 106, venter of idiosoma; Fig. 107, dorsum of
idiosoma and gnathosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 108 – 112
Unionicola sp. (Unionicolinae), larva. Fig. 108, Dorsum of idiosoma and gnathosoma;
Fig. 109, excretory pore plate; Fig. 110, claws and empodium of leg I; Fig. 111, venter of idiosoma and
gnathosoma; Fig. 112, tibia and tarsus of leg I. FIGURE 113 Koenikea marshallae Viets (Pionatacinae),
larva, excretory pore plate. FIGURES 114 – 116 Neumania punctata Marshall (Pionatacinae), larva. Fig.
114, excretory pore plate; Fig. 115, claws and empodium of leg I; Fig. 116, tibia and tarsus of leg I.
595
596
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 117 – 118 Wettina ontario Smith (Wettinidae), larva. Fig. 117, Venter of idiosoma; Fig. 118,
excretory pore plate. FIGURES 119 – 120 Hydrochoreutes intermedius Cook (Hydrochoreutinae), larva.
Fig. 119, Excretory pore plate; Fig. 120, venter of idiosoma and gnathosoma. FIGURE 121 Hydrochoreutes
minor Cook (Hydrochoreutinae), larva, excretory pore plate. FIGURES 122 – 123 Huitfeldtia rectipes
Thor (Huitfeldtiinae), larva. Fig. 122, Venter of idiosoma; Fig. 123, excretory pore plate. FIGURES 124 – 125
Neotiphys pionoidellus (Habeeb) (Tiphyinae), larva. Fig. 124, Venter of idiosoma; Fig. 125, excretory pore plate.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 126 – 127
Tiphys americanus (Marshall) (Tiphyinae), larva. Fig. 126, Venter of idiosoma and
gnathosoma; Fig. 127, excretory pore plate. FIGURES 128, 130 Tiphys ornatus (Koch) (Tiphyinae), larva.
Fig. 128, Tibia and tarsus of leg I; Fig. 130, excretory pore plate. FIGURES 129, 131 Pseudofeltria
multipora Cook (Foreliinae), larva. Fig. 129, Venter of idiosoma; Fig. 131, trochanter of leg III. FIGURES
132 – 134 Forelia ovalis Marshall (Foreliinae), larva. Fig. 132, Tibia and tarsus of leg III; Fig. 133, venter of
idiosoma and gnathosoma; Fig. 134, excretory pore plate. FIGURES 135, 136 Forelia onondaga Habeeb
(Foreliinae), larva. F ig. 135, excretory pore plate; Fig. 136, venter of idiosoma.
597
598
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 137 – 138, 141 Najadicola ingens Koenike (Najadicolinae), larva. Fig. 137, Venter of idiosoma;
Fig. 138, excretory pore plate; Fig. 141, tibia and tarsus of leg II. FIGURES 139, 140 Nautarachna muskoka
Smith (Pioninae), larva. Fig. 139, Excretory pore plate; Fig. 140, venter of idiosoma. FIGURES 142, 143
Piona carnea (Koch) (Pioninae), larva. Fig. 142, excretory pore plate; Fig. 143, claws and empodium of leg III.
FIGURES 144, 145, 148 Piona interrupta Marshall (Pioninae), larva. Fig. 144, dorsum of idiosoma and
gnathosoma; Fig. 145, venter of idiosoma and gnathosoma; Fig. 148, excretory pore plate. FIGURE 146
Piona constricta (Wolcott) (Pioninae), larva, excretory pore plate. FIGURES 147, 149 Piona mitchelli Cook
(Pioninae), larva. Fig. 147, tarsus of leg I; Fig. 149, tibia and tarsus of leg II.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 150 – 152 Ljania bipapillata Thor (Axonopsinae), larva. Fig. 150, Dorsum of idiosoma and
gnathosoma; Fig. 151, tibia and tarsus of leg I; Fig. 152, venter of idiosoma. FIGURES 153–154 Albia
neogaea Habeeb (Albiinae), larva. Fig. 153, Dorsum of idiosoma and gnathosoma; Fig. 154, venter of
idiosoma. FIGURE 155 Neobrachypoda ekmani (Walter) (Axonopsinae), larva, venter of idiosoma.
599
600
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 156,160 Woolastookia setosipes Habeeb (Axonopsinae), larva. Fig. 156, Venter of idiosoma;
Fig. 160, excretory pore plate. FIGURES 157–159 Estellacarus unguitarsus (Habeeb) (Axonopsinae), larva.
Fig. 157, Venter of idiosoma, Fig. 158, tibia and tarsus of leg I; Fig. 159, excretory pore plate. FIGURE 161
Woolastookia pilositarsa (Habeeb) (Axonopsinae), larva, excretory pore plate. FIGURE 162 Brachypoda
cornipes Habeeb (Axonopsinae), larva, excretory pore plate. FIGURE 163 Brachypoda setosicauda Habeeb
(Axonopsinae), larva, excretory pore plate.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 164 Brachypoda setosicauda Habeeb (Axonopsinae), larva, venter of idiosoma. FIGURES
165–167 Axonopsis setoniensis Habeeb (Axonopsinae), larva. Fig. 165, venter of idiosoma; Fig. 166,
excretory pore plate; Fig. 167, tibia and tarsus of leg I. FIGURES 168–170 Aturus sp. (Aturinae), larva.
Fig. 168, dorsum of idiosoma and gnathosoma; Fig. 169, venter of idiosoma; Fig. 170, tibia and tarsus of leg I.
601
602
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 171 – 172, 175 Stygomomonia mitchelli Smith (Stygomomoniinae), larva. Fig. 171, dorsum of
idiosoma and gnathosoma; Fig. 172, venter of idiosoma; Fig. 175, distal segments of leg III. FIGURE 173
Momonia campylotibia Smith (Momoniinae), larva, distal segments of leg III. FIGURES 174, 177
Paramideopsis susanae Smith (Nudomideopsidae), larva. Fig. 174, tibia and tarsus of leg III; Fig. 177,
pedipalp. FIGURE 176 Nudomideopsis magnacetabula (Smith), larva, excretory pore plate.
16. Water Mites (Hydrachnida) and Other Arachnids
FFIGURES 178 – 179 Nudomideopsis magnacetabula (Smith) (Nudomideopsidae), larva. Fig. 178, dorsum
of idiosoma and gnathosoma; Fig. 179, venter of idiosoma. FIGURES 180–181, 183 Midea expansa
Marshall (Mideidae), larva. Fig. 180, venter of idiosoma; Fig. 181, tibia and tarsus of leg III; Fig. 183, venter
of gnathosoma. FIGURES 182, 184–185 Acalyptonotus neoviolaceus Smith (Acalyptonotidae), larva.
Fig. 182, pedipalp; Fig. 184, dorsum of idiosoma and gnathosoma; Fig. 185, venter of idiosoma.
603
604
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 186, 188 Mideopsis borealis Habeeb (Mideopsinae), larva. Fig. 186, venter of idiosoma; Fig. 188,
dorsum of idiosoma and gnathosoma. Figures 187, 189–190 Platyhydracarus juliani Smith (Athienemanniinae), larva. Fig. 187, tibia and tarsus of leg II; Fig. 189, venter of idiosoma; Fig. 190, dorsum of idiosoma and
gnathosoma. FIGURE 191 Neoacarus occidentalis Cook (Neoacaridae), larva, dorsum of idiosoma and
gnathosoma. FIGURE 192 Laversia berulophila Cook (Larversiidae), larva, venter of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 193, 195 – 196 Arrenurus planus Marshall (Arrenurinae), larva. Fig. 193, venter of idiosoma;
Fig. 195, dorsum of idiosoma and gnathosoma; Fig. 196, tibia and tarsus of leg III. FIGURE 194 Laversia
berulophila Cook (Laversiidae), larva, tibia and tarsus of leg II. FIGURES 197–198 Krendowskia similis
Viets (Krendowskiidae), larva. Fig. 197, venter of idiosoma; Fig. 198, tibia and tarsus of leg III.
605
606
Ian M. Smith, David R. Cook, Bruce P. Smith
2. Taxonomic Key to Genera for Adult Water Mites
1a.
Legs with tarsi bearing empodium and paired claws (Fig. 201); pedipalps with tarsus bearing single, long terminal seta (Fig. 202,
arrow); idiosoma soft and elongate.......................................................Stygothrombidioidea Stygothrombidiidae Stygothrombium
1b.
Legs with tarsi bearing paired claws, but lacking empodium; pedipalps with tarsus lacking single long terminal seta as above;
idiosoma various ............................................................................................................................................................................2
2a(1b).
Genital field with movable genital flaps, but with genital acetabula borne on coxal plates (Fig. 205, arrow) ......Hydrovolzioidea 3
2b.
Genital field with or without movable genital flaps; when genital flaps present, genital acetabula borne on them (Fig. 200) or beneath them (Fig. 283) .....................................................................................................................................................................4
3a(2a).
Dorsal shield with large central plate surrounded by smaller unpaired plate anteriorly and numerous pairs of small similar
platelets laterally and posteriorly (Fig. 204); glandularia represented by setae, with gland portions absent ......................................
....................................................................................................................................................Acherontacaridae Acherontacarus
3b.
Dorsal shield with large central plate surrounded by smaller unpaired plate anteriorly and several pairs of small disimilar plates
laterally and posteriorly (Fig. 206); glandularia divided into gland and seta bearing sclerites (Fig. 203)...........................................
............................................................................................................................................................Hydrovolziidae Hydrovolzia
4a(2b).
Idiosoma nearly spherical (Fig. 207); pedipalps with tibia much shorter than genu and bearing a distidorsal projection extending
well beyond insertion of tarsus (Fig. 209); chelicerae 1-segmented (Fig. 208)...................................................................................
.......................................................................................................................................Hydrachnoidea Hydrachnidae Hydrachna
4b.
Idiosoma variously shaped, but rarely spherical; pedipalps with tibia usually longer than genu, when shorter then lacking a
distidorsal projection; chelicera 2-segmented (Fig. 316) .................................................................................................................5
5a(4b).
Lateral eye capsules borne on medial prodorsal plate (Figs. 210, 211, 212) or, in one family, on moderately large, rugose, paired
platelets associated with complex of six prodorsal sclerites (Fig. 213)............................................................................Eylaoidea 6
5b.
Lateral eye capsules, when present, relatively small and not borne on prodorsal plate (Figs. 220, 224)........................................11
6a(5a).
Dorsum with lateral eyes borne on lateral platelets separated by a medial triangular platelet (Fig. 213, arrows); idiosoma bearing
26 pairs of glandularia on large, rugose platelets (Figs. 213, 214); gnathosoma with capitulum wider than long and pedipalps
with only one movable segment which flexes medially to oppose anetrior edge of capitulum (Fig. 214)...........................................
.............................................................................................................................Apheviderulicidae Apheviderulix (deutonymphs)
6b.
Dorsum with lateral eyes borne on a medial prodorsal plate (Fig. 210, 211, 212); idiosoma bearing fewer than 26 pairs of glandularia on small sclerites; gnathosoma with capitulum longer than wide and pedipalps with 3 to 5 movable segments which flex
ventrally .........................................................................................................................................................................................7
7a(6b).
Lenses of lateral eyes well separated from each other on their respective sides and borne laterally on entire prodorsal plate bearing one pair of setae (Fig. 210) .................................................................................................................................Eylaidae Eylais
7b.
Lenses of lateral eyes relatively close together on their respective sides and borne laterally on smooth, entire prodorsal plate bearing four pairs of setae (Figs. 211, 212) ...........................................................................................................................................8
8a(7b).
Prodorsal plate approximately as wide as long (Fig. 212); genital acetabula surrounded by acetabular plates..................................
.........................................................................................................................................................................Piersigiidae Piersigia
8b.
Prodorsal plate much longer than wide (Fig. 211); genital acetabula scattered in ventral integument ....................Limnocharidae 9
9a(8b).
Gnathosoma protrusible; pedipalps three segmented, with tarsus very small and inserted dorsally on tibia (Fig. 215, arrow)
...................................................................................................................................Rhyncholimnocharinae Rhyncholimnochares
9b.
Gnathosoma not protrusible; pedipalps 4- or 5-segmented, with tarsus inserted at distal end of tibia (Figs. 216, 217) ....................
............................................................................................................................................................................Limnocharinae 10
10a(9b).
Pedipalps 4-segmented (Fig. 217)............................................................................................................................Neolimnochares
10b.
Pedipalps 5-segmented (Fig. 216) .................................................................................................................................Limnochares
11a(5b).
Dorsum with a series of reticulate platelets as shown in Figure 218 ................................................................................................
..........................................................................................................Rhynchohydracaridae Clathrosperchoninae Clathrosperchon
11b.
Dorsum various, but not with an arrangement of platelets as shown in Figure 218......................................................................12
12a(11b).
Pedipalps chelate, with dorsodistal portion of tibia extending well beyond insertion of tarsus (Figs. 221, 227, arrows); capitulum
without anchoral process (as in Fig. 342, arrow)..........................................................................................................................13
12b.
Pedipalps not chelate, when appearing chelate (as in some Tiphyinae) (Fig. 332) then capitulum with well-developed anchoral
process (Fig. 330, arrow A) ..........................................................................................................................................................38
13a(12a).
Pedipalps with dorsodistal extension of tibia relatively long (Fig. 221, arrow)...................................Hydrodromidae Hydrodroma
16. Water Mites (Hydrachnida) and Other Arachnids
607
13b.
Pedipalps with dorsodistal extension of tibia relatively short (Fig. 227, arrow) .................................................Hydryphantidae 14
14a(13b)
Legs with swimming setae on distal segments...............................................................................................................................15
14b.
Legs without swimming setae ......................................................................................................................................................16
15a(14a).
Dorsum with medial eye plate between lateral eyes bearing a pigmented medial eye and two pairs of setae (Figs. 219, 220) ...........
........................................................................................................................................................Hydryphantinae Hydryphantes
15b.
Dorsum without medial eye plate between lateral eyes ................................................Pseudohydryphantinae Pseudohydryphantes
16a(14b).
Third and fourth coxal plates with medial margins extensive and close together, not separated by genital field (Fig. 223); lateral
eyes in capsules; body not elongated......................................................................................................Cowichaniinae Cowichania
16b.
Posterior coxal groups usually well separated from each other, with genital field extending between them (Fig. 237), when these
coxal plates close together then lateral eyes not in capsules and body greatly elongated (Fig. 226) ..............................................17
17a(16b).
Lateral eyes in capsules (Fig. 224, arrow).....................................................................................................................................18
17b.
Lateral eyes, when present, not in capsules...................................................................................................................................37
18a(17a).
Dorsum without dorsalia (Figs. 224, 230)....................................................................................................................................19
18b.
Dorsum with dorsalia of various sizes and arrangement (Figs. 232, 234, 245, 256) .....................................................................22
19a(18a).
Genital field bearing three pairs of acetabula ................................................................................................Thyadinae (in part) 20
19b.
Genital field bearing numerous acetabula (Fig. 228) ....................................................................................................Protziinae 21
20a(19a)
Dorsum bearing well-developed medial eye containing two dots of pigment (Fig. 225); genital field with setiferous extensions of
genital flaps extending anterior to first pair of acetabula (Fig. 229, arrow)...................................................................Notopanisus
20b.
Dorsum lacking medial eye (Fig. 230); genital field without setiferous extensions of genital flaps extending anterior to first pair of
acetabula (Fig. 231)......................................................................................................................................................Albertathyas
21a(19b).
Legs with tarsal claws bearing palmately arranged clawlets (Fig. 222) ..................................................................................Protzia
21b.
Legs with tarsal claws simple ............................................................................................................................................Partnunia
22a(18b).
Dorsum with all dorsalia peripheral in position (Fig. 232, arrow) .........................................................Cyclothyadinae Cyclothyas
22b.
Dorsum with some dorsalia more medial in position (Figs. 245, 256)...........................................................Thyadinae (in part) 23
23a(22b).
Dorsum with single, large anterior plate, but without other dorsalia (Fig. 233) ..........................................................Siskiyouthyas
23b.
Dorsum with or without single, large anterior plate, but always bearing other paired dorsalia ....................................................24
24a(23b).
Dorsum with large anterior plate, and with two medial and four lateral pairs of platelets (Fig. 234).............................Trichothyas
24b.
Dorsum with dorsalia variously developed and fused, but not as above .......................................................................................25
25a(24b).
Dorsum with medial eye borne on spindle-shaped sclerite (Fig. 235, arrow); genital field with second pair of acetabula located
about halfway between first and third pairs (Figs. 236, 237) ...............................................................................................Euthyas
25b.
Dorsum with medial eye not borne on spindle-shaped sclerite; genital field with second pair of acetabula usually located in posterior half of gonopore ...................................................................................................................................................................26
26a(25b).
Dorsum with dorsalia fused into single dorsal shield (i.e., unsclerotized integument surrounds dorsoglandularia, but does not occur between dorsalia) (Figs. 238, 240, 241)..................................................................................................................................27
26b.
Dorsum with dorsalia not fused into single dorsal shield (i.e., unsclerotized integument separates dorsalia) (Figs. 242, 245).......29
27a(26a).
Genital field with flaps not extending anterior to first pair of acetabula, and second pair of acetabula located beside third pair
(Fig. 239) ..........................................................................................................................................................................Thyopsis
27b.
Genital field with flaps extending anterior to first pair of acetabula, and second pair of acetabula located well anterior to third
pair (Figs. 244, 246).....................................................................................................................................................................28
28a(27b).
Dorsal shield well sclerotized, with some of thickened areas indicating medial dorsalia and adjacent lateral dorsalia fused on each
side (Fig. 240, arrows) (edges of dorsalia obscured by secondary sclerotization in mature specimens); genital field with setiferous
projections of genital flaps medial to third pair of acetabula relatively small (Fig. 244) ..................................................Thyopsella
28b.
Dorsal shield weakly sclerotized, with thickened areas indicating medial and lateral dorsalia separate (Fig. 241); genital field with
setiferous projections of genital flaps medial to third pair of acetabula relatively large (Fig. 246).................................Thyopsoides
29a(26b).
Dorsum with most posterior pair of dorsalia fused into single large plate (Figs. 242, 243, arrows)..............................................30
29b.
Dorsum with most posterior pair of dorsalia separated medially (Figs. 245, 248, 250) ................................................................31
608
Ian M. Smith, David R. Cook, Bruce P. Smith
30a(29a).
Dorsum with central region bearing three pairs of dorsalia that are separated medially (Fig. 242) .......................................Panisus
30b.
Dorsum with central region bearing two pairs of dorsalia that are fused medially (Fig. 243)....................................Columbiathyas
31a(29b).
Genital field with well sclerotized extensions of genital flaps extending anterior to first pair of acetabula (Figs. 247, 249,
arrows).........................................................................................................................................................................................32
31b.
Genital field without sclerotized extensions of genital flaps anterior to first pair of acetabula (Fig. 253) ......................................34
32a(31a).
Dorsum with lateral eye capsules raised and located dorsally on idiosoma (Fig. 245, arrow A) and with medial eye well
developed (Fig. 245, arrow B); genital field with setae on anterior extensions of genital flaps relatively thick (Fig. 247, arrow)
........................................................................................................................................................................................Panisopsis
32b.
Dorsum with lateral eye capsules relatively flat and located at anterior edge of idiosoma (Fig. 248, arrow) and with medial
eye indistinct or absent (Figs. 248); genital field with setae on anterior extension of genital flaps relatively slender (Fig. 249,
arrow) ..........................................................................................................................................................................................33
33a(32b).
Dorsum with four pairs of medial dorsalia posterior to anteromedial plate (Fig. 248).................................................Parathyasella
33b.
Dorsum with five pairs of medial dorsalia posterior to anteromedial plate (Fig. 251)........................................................Thyasella
34a(31b).
Dorsum with dorsalia large and occupying most of the dorsal area of the idiosoma (Fig. 250) ................................Amerothyasella
34b.
Dorsum with dorsalia small and occupying only a small part of the dorsal area of the idiosoma (Figs. 252, 256)........................35
35a(34b).
Dorsum with medial eye bearing two pigment dots and usually borne on small platelet formed by fused pre — and postocularia
(Fig. 252, arrow)...............................................................................................................................................................Thyasides
35b.
Dorsum with medial eye unpigmented or with dispersed pigment, and not borne on small platelet (Fig. 256) .............................35
36a(35b).
Genital field with posterior two pairs of acetabula not incorporated into edges of genital flaps (Fig. 253)..............................Thyas
36b.
Genital field with posterior two pairs of acetabula fused with edges of genital flaps (Fig. 257) .......................................Zschokkea
37a(17b).
Body not greatly elongated; genital flaps well developed (Fig. 254, arrow).......................................Tartarothyadinae Tartarothyas
37b.
Body greatly elongated (Fig. 226); genital flaps poorly developed or absent..................................................Wandesiinae Wandesia
38a(12b).
Femur of pedipalp with two long medial setae (Fig. 255, arrow); occuring only in hot springs ...........Thermacaridae Thermacarus
38b.
Femur of pedipalp usually without two medial setae and never as illustrated in Figure 255; not occurring in hot springs............39
39a(38b).
Genital field with movable genital flaps flanking gonopore and either partially or completely covering gonopore when closed;
genital acetabula lying free in gonopore (not on flaps or acetabular plates) (Figs. 259, 277, 283) ...........................Lebertioidea 40
39b.
Genital field usually without movable genital flaps, when flaps present then acetabula lie on flaps rather than in gonopore
(Fig. 200) .....................................................................................................................................................................................59
40a(39a).
Fourth coxal plates with medial edges reduced to narrow angles and bearing a pair of glandularia (Fig. 264, arrow); tarsus of
pedipalp bearing pad-shaped setae and appearing spatulate distally (Fig. 263, arrow) ............................Rutripalpidae Rutripalpus
40b.
Fourth coxal plates with medial edges usually extensive, when reduced to narrow angles then lacking glandularia as above.......41
41a(40b).
Dorsal and ventral shields present; dorsal shield usually comprising a large plate and a series of smaller platelets (Figs. 273, 276,
278); genital field with gonopore usually bearing six pairs of acetabula (Fig. 279), when only three pairs of genital acetabula present then dorsal shield with complete ring of marginal platelets (Fig. 273); one known North American species has platelets fused
into an entire dorsal shield, but six pairs of genital acetabula are diagnostic .......................................................Torrenticolidae 42
41b.
Dorsal and ventral shields present or absent; when present, dorsal shield entire, without smaller platelets; genital field with gonopore bearing three pairs of acetabula............................................................................................................................................46
42a(41a).
Dorsal shield with single anteromedial platelet and series of small marginal platelets (Fig. 273); genital field with gonopore bearing three pairs of genital acetabula ......................................................................................................Testudacarinae Testudacarus
42b.
Dorsal shield with one or two pairs of anterior platelets (Figs. 276, 278) (in one species platelets are fused with large central
plate); genital field with gonopore bearing six pairs of acetabula.........................................................................Torrenticolinae 43
43a(41b).
Pedipalps 4-segmented (Fig. 275)................................................................................................................................Neoatractides
43b.
Pedipalps 5-segmented (as in Fig. 274) ........................................................................................................................................44
44a(43b).
Gnathosoma protrusible, capitulum long and slender(Fig. 280)..........................................................................Pseudotorrenticola
44b.
Gnathosoma not protrusible, capitulum relatively short and stocky (Fig. 281).............................................................................45
45a(44b).
Capitulum with short proximodorsal projections (Fig. 281, arrow) .............................................................................Torrenticola
45b.
Capitulum with long proximodorsal projections (Fig. 282, arrow) .........................................................................Monoatractides
16. Water Mites (Hydrachnida) and Other Arachnids
609
46a(41b).
Fourth coxal plates bearing glandularia (Fig. 265, arrow)...............................................................................Teutoniidae Teutonia
46b.
Fourth coxal plates lacking glandularia (Figs. 271, 283) ..............................................................................................................47
47a(46b).
Venter without medial suture line separating coxal plates (Fig. 289); all legs inserted near anterior end of idiosoma (Fig. 289)
........................................................................................................................................................................................Oxidae 48
47b.
Venter with medial suture line separating coxal plates of either side (Fig. 283, arrow A); fourth pair of legs inserted laterally near
middle of idiosoma (Fig. 283, arrow C) .......................................................................................................................................49
48a(47a).
Idiosoma strongly compressed laterally with coxal plates extending onto dorsal surface leaving only narrow medial unsclerotized
strip of soft integument that usually bears small sclerites (Fig. 291); sexual dimorphism not pronounced ......................Frontipoda
48b.
Idiosoma ovoid or moderately compressed laterally, without small sclerites in dorsal integument (Fig. 288); sexual dimorphism
usually not pronounced, but in some species coxal plates extend more dorsally in males .......................................................Oxus
49a(47b).
Venter with suture lines between second and third coxal plates incomplete (Fig. 283, arrow B), but touching the genital field area
posteriorly; genu of pedipalp usually with at least five long setae on medial surface (Fig. 284), but, in one known species, with
four long setae on dorsal surface ...............................................................................................................................Lebertiidae 50
49b.
Venter with suture lines between second and third coxal plates complete (Fig. 268) and these coxal plates often completely
separated on their respective sides (Fig. 259); genu of pedipalp without long setae ......................................................................52
50a(49a).
Idiosoma strongly compressed laterally (Fig. 285)............................................................................................................Estelloxus
50b.
Idiosoma ovoid or dorsoventrally flattened ..................................................................................................................................51
51a(50b).
Dorsum covered by complete dorsal shield (Fig. 286); pedipalps with femur bearing several long setae dorsally and genu bearing
four long setae dorsally (Fig. 287)................................................................................................................................Scutolebertia
51b.
Dorsum usually covered by soft integument, rarely covered by dorsal shield; pedipalps with femur bearing few short setae and
genu bearing five or six long setae on medial surface (Fig. 284) ..........................................................................................Lebertia
52a(49b).
Dorsum and venter covered by complete shields...................................................................................................Anisitsiellidae 53
52b.
Dorsum and venter with sclerites variously developed, but complete dorsal and ventral shields never both present .........................
..............................................................................................................................................................................Sperchontidae 58
53a(52a).
Tarsi of fourth pair of legs lacking claws (Fig. 267)....................................................................................................Mamersellides
53b.
Tarsi of fourth pair of legs bearing claws (Fig. 266) .....................................................................................................................54
54a(53b).
Fourth pair of legs inserted at anterolateral angles of fourth coxal plates, just posterior to insertions of third pair of legs (Fig.
268, arrow) ......................................................................................................................................................................Utaxatax
54b.
Fourth pair of legs inserted near posterolateral angles of fourth coxal plates, well posterior to insertions of third pair of legs
(Fig. 271, arrow A) ......................................................................................................................................................................55
55a(54b).
Accessory glandularia (“Glandula Limnesiae”) located in concavities of anteromedial edges of fourth coxal plates (Fig. 269,
arrow) .........................................................................................................................................................................Bandakiopsis
55b.
Accessory glandularia located on first or third coxal plates ..........................................................................................................56
56a(55b).
Accessory glandularia located near anterior angles of first coxal plates (Fig. 270, arrow)..............................................Cookacarus
56b.
Accessory glandularia located on third coxal plates .....................................................................................................................57
57a(56b).
Accessory glandularia located near medial edges of third coxal plates (Fig. 271, arrow B); fourth coxal plates without or with
only small projections associated with insertions of fourth pair of legs..............................................................................Bandakia
57b.
Accessory glandularia located near lateral edges of third coxal plates (Fig. 272, arrow A); fourth coxal plates with large projections associated with insertions of fourth pair of legs (Fig. 272, arrow B) ..................................................................Oregonacarus
58a(52b).
Idiosoma with glandularia borne on smooth platelets that are usually small and flat, rarely enlarged and raised (Fig. 258);
pedipalps with tibia usually bearing two peglike setae ventrally (Fig. 260, arrows)............................................................Sperchon
58b.
Idiosoma with glandularia borne on tuberculate or papillate platelets that are enlarged and raised (Fig. 261); pedipalps with tibia
lacking peg-like setae ventrally (Fig. 262)...................................................................................................................Sperchonopsis
59a(39b).
All coxal plates grouped together, with medial edges of anterior coxal groups noticeably longer than those of posterior coxal
groups (Fig. 292); fourth coxal plates rounded posteriorly and suture lines between third and fourth plates extending posterolaterally (Fig. 292)...................................................................................................................................Omartacaridae Omartacarus
610
Ian M. Smith, David R. Cook, Bruce P. Smith
59b.
Coxal plates not as described and illustrated above......................................................................................................................60
60a(59b).
Pedipalps with femur usually bearing single ventral seta (Fig. 296, 306, arrows), when this seta absent then tarsi of pedipalps
with curved terminal seta (Fig. 297, arrow); genital field with movable genital flaps which cover gonopore when closed in females
(Fig. 200) and some males ........................................................................................................................................Limnesiidae 61
60b.
Pedipalps with femur lacking ventral setae or, rarely, bearing three setae (Fig. 414, arrow); genital field without movable genital
flaps which cover gonopore when closed......................................................................................................................................69
61a(60a).
Pedipalps with tibia bearing five or more long, pointed ventral tubercles and a long medial seta (Fig. 296); idiosoma lacking dorsal and ventral shields, although dorsal plates present ...............................................................Kawamuracarinae Kawamuracarus
61b.
Pedipalps with tibia bearing four long, pointed tubercles (Fig. 295) or lacking tubercles (Fig. 298), and lacking a long medial
seta; idiosoma with dorsal and ventral shields..............................................................................................................................62
62a(61b).
Fourth pair of legs with tarsus lacking claws................................................................................................................................63
62b.
Fourth pair of legs with tarsus bearing claws ...............................................................................................................................67
63a(62a).
Genital field with genital flaps on each side of gonopore divided into anterior and posterior sclerites (Fig. 299); dorsum with
large, transversely divided dorsal shield (Fig. 294) ............................................................................................Neomamersiinae 64
63b.
Genital field with genital flaps of females (and acetabular plates of males) entire on each side (Fig. 200); dorsum usually without
dorsal shield, but rarely with entire (undivided) shield..............................................................................................Limnesiinae 66
64a(63a).
Ventral shield with sclerotized bridge forming dorsal side of camerostome incomplete (Fig. 304, arrow A); venter with posterior
region bearing several small platelets (Fig. 304, arrow B) or at least a small excretory pore platelet separate from ventral shield;
genital field of both sexes with middle pair or group of acetabula borne on posterior sclerites of genital flaps (Fig. 300); pedipalps
with femur and tibia relatively long and slender and tibia lacking ventral tubercles (Fig. 298)........................................Meramecia
64b.
Ventral shield with sclerotized bridge forming dorsal side of camerostome complete (Fig. 302, arrow A); ventral shield entire posteriorly (Figs. 302, 303); genital field of males with middle pair or group of acetabula borne on anterior sclerites of genital flaps
(Fig. 301), or on pair of separate, small, medial sclerites (Fig. 299); pedipalps not as above ........................................................65
65a(64b).
Pedipalps with femur bearing ventral seta, tibia bearing four long ventral tubercles and tarsus lacking long, curved distal seta
(Fig. 295).....................................................................................................................................................................Neomamersa
65b.
Pedipalps with femur lacking ventral seta, tibia lacking ventral tubercles and tarsus bearing long, curved distal seta (Fig. 297,
arrow) .........................................................................................................................................................................Arizonacarus
66a(65b).
Genital field of males immediately flanked by pair of glandularia (Fig. 305).............................................................Centrolimnesia
66b.
Genital field of males not immediately flanked by pair of glandularia ................................................................................Limnesia
67a(62b).
Dorsum nearly covered by two large plates (Fig. 307); pedipalps with femur bearing ventral seta inserted directly on surface of
segment (as in Fig. 296, arrow)........................................................................................................Protolimnesiinae Protolimnesia
67b.
Dorsum entirely covered by dorsal shield (Fig. 310) or less than half covered by small platelets (Fig. 311); pedipalps with femur
bearing ventral seta inserted on prominent tubercle (Fig. 306, arrow) .......................................................................Tyrrelliinae 68
68a(67b).
Third coxal plates with pair of accessory glandularia (“Glandula Limnesiae”) (Fig. 308, arrow); genital field with numerous
small acetabula (Fig. 308)..............................................................................................................................................Neotyrrellia
68b.
Third coxal plates lacking glandularia (Fig. 309); genital field with 3 – 7 pairs of relatively large acetabula (Fig. 309) ........Tyrrellia
69a(60b).
Integument between fourth coxal plates and genital field with two pairs of glandularia arranged more or less in a row, and either
free (Fig. 312, arrows) or variously fused with other ventral sclerites; dorsal shield absent, or present and bearing fewer than five
pairs of glandularia.................................................................................................................................................Feltriidae Feltria
69b.
Integument between fourth coxal plates and genital field usually without two pairs of glandularia arranged in a row, when glandularia in this position (some Koenikea) then dorsal shield bearing six pairs of glandularia (Fig. 322) ........................................70
70a(69b).
Idiosoma strongly compressed laterally and much higher than wide (Fig. 314); fourth legs with all segments flattened (Fig. 313)
.....................................................................................................................................................Frontipodopsidae Frontipodopsis
70b.
Idiosoma not strongly compressed laterally..................................................................................................................................71
71a(70b).
Both dorsal and ventral shields present; first pair of legs with tibia lacking two thickened setae and a downturned seta
distoventrally (ie. not as shown in Fig. 321) ................................................................................................................................72
71b.
Either dorsal or ventral shield, or both, usually absent, when both shields present (males of some Atractides) then first pair of
legs with tarsus bowed and tibia bearing two thickened setae and a downturned seta distoventrally (Fig. 321) .........................125
16. Water Mites (Hydrachnida) and Other Arachnids
611
72a(71a).
First leg with tarsus much shorter than tibia, and claws dorsal in position and flexing proximally (Fig. 381, arrow) .......................
................................................................................................................................................................................Momoniidae 73
72b.
First leg with distal segments not modified as described and illustrated above; claws terminal in position and flexing ventrally...75
73a(72a).
Dorsal shield composed of large plate surrounded by numerous smaller platelets (Fig. 380) ......Cyclomomoniinae Cyclomomonia
73b.
Dorsal shield entire or comprising large anterior and posterior plates, but not surrounded by smaller platelets (ie. not as in Fig.
380) .............................................................................................................................................................................................74
74a(73b).
Venter with pair of glandularia located close to anterior end of genital field (Fig. 382, arrow); genital field of males with acetabula borne in gonopore..................................................................................................................................Momoniinae Momonia
74b.
Venter without glandularia near anterior end of genital field (Fig. 383); genital field of males with acetabula borne on plates
flanking gonopore (Fig. 383)........................................................................................................Stygomomoniinae Stygomomonia
75a(72b).
Dorsal shield large and transversely divided (somewhat as in Fig. 294); suture lines between third and fourth coxal plates noticeably looped around a pair of glandularia (Fig. 293, arrow) ................................................Hygrobatidae (in part) Diamphidaxona
75b.
Dorsal shield usually entire, never transversely divided as above; suture lines between third and fourth coxal plates not looped as
above ...........................................................................................................................................................................................76
76a(75b).
Venter with outer edges of coxal plates forming a smooth arc which does not extend beyond edge of idiosoma (Fig. 389, arrow);
pedipalps highly modified (Fig. 390)...............................................Chappuisididae (in part) Uchidastygacarinae Uchidastygacarus
76b.
Venter with outer edges of coxal plates not forming a smooth arc; pedipalps not as described and illustrated above ...................77
77a(76b).
Venter with pair of glandularia located near midline at junction of third and fourth coxal plates (Fig. 388, arrow).........................
....................................................................................................................Chappuisididae (in part) Chappuisidinae Chappuisides
77b.
Venter without glandularia in position described and illustrated above ........................................................................................78
78a(77b).
Ventral shield with suture lines between third and fourth coxal plates oblique, extending posteromedially to genital field region
and well separated from each other medially (Fig. 395, arrow); genital acetabula borne in single rows on each side, either in
gonopore (males) or on slender plates flanking gonopore (females) (Fig. 395) .........................................................Neoacaridae 79
78b.
Ventral shield not with above combination of characters .............................................................................................................80
79a(78a).
Pedipalps with tibia moderately produced distoventrally to oppose tarsus (Fig. 396) ......................................................Neoacarus
79b.
Pedipalps with tibia and tarsus highly modified to form chelate appendage (Fig. 397) .................................................Volsellacarus
80a(78b).
Ventral shield with coxoglandularia I shifted far forward in coxal plates II, near suture lines between first and second coxal plates
(Figs. 384 – 387, arrows)...............................................................................................................................................................81
80b.
Ventral shield with coxoglandularia I located between second and third coxal plates (Fig. 407, arrow) or in posterior edges of
second coxal plates (Fig. 406, arrow A) .......................................................................................................................................84
81a(80a).
Ventral shield with genital field projecting between, and widely separating, fourth coxal plates (Fig. 384) .............Mideidae Midea
81b.
Ventral shield with genital field located posterior to fourth coxal plates (Fig. 386) .........................................Nudomideopsidae 82
82a(81b).
Genital field with three pairs of acetabula (Fig. 385)................................................................................................Nudomideopsis
82b.
Genital field with more than three pairs of acetabula ...................................................................................................................83
83a(82b).
Genital field with four-to-six pairs of acetabula arranged in single rows (Fig. 386)....................................................Paramideopsis
83b.
Genital field with more than six pairs of acetabula arranged in several rows (Fig. 387) .............................................Neomideopsis
84a(80b).
Pedipalps distinctly uncate (Fig. 401; Fig. 399, arrow A; Fig. 416, arrow) ...................................................................................85
84b.
Pedipalps not distinctly uncate, with tibia bearing either slight distoventral projection (Fig. 417, arrow), or distinct ventral projection near middle of segment (Fig. 405), or no ventral projection..............................................................................................91
85a(84a).
Ventral shield with fourth coxal plates bearing pair of glandularia (Fig. 413, arrow); genital field extending between fourth coxal
plates, widely separating them medially, and usually bearing three or four pairs of genital acetabula (but with five or six pairs in a
few species) ........................................................................................................................................................Krendowskiidae 86
85b.
Ventral shield with fourth coxal plates lacking glandularia; genital field extending at most only slightly between fourth coxal
plates, not separating them medially, and bearing numerous acetabula (Fig. 402) ........................................................................87
86a(85a).
Gnathosoma protrusible........................................................................................................................................................Geayia
612
Ian M. Smith, David R. Cook, Bruce P. Smith
86b.
Gnathosoma not protrusible........................................................................................................................................Krendowskia
87a(85b).
Genital field with some or all acetabula lying free in gonopore (Fig. 402), or with acetabula lying on plates which flank gonopore, but are not fused with ventral shield (similar to Fig. 384); capitulum with a pair of long ventral setae (Fig. 401, arrow);
tibia of pedipalp rotated relative to genu (up to 90°).......................................................................................Athienemanniidae 88
87b.
Genital field with acetabula borne on acetabular plates incorporated into ventral shield, never in gonopore (Fig. 415); capitulum
with all setae short; tibia of pedipalp not greatly rotated relative to genu (Fig. 416); posterior end of idiosoma variously (and often highly) modified in males (Figs. 419, 420)...........................................................................Arrenuridae Arrenurinae Arrenurus
88a(87a).
Pedipalp with genu bearing pronounced ventral projection (Fig. 399, arrow B); genital field with acetabula in one or two rows on
each side of gonopore (Fig. 398)..................................................................................................Stygameracarinae Stygameracarus
88b.
Pedipalp with genu lacking pronounced ventral projection (Figs. 401); genital field with acetabula in one to many rows on each
side of gonopore (Figs. 402 – 404) ...................................................................................................................Athienemanniinae 89
89a(88b).
Venter with coxal plates I extending anteriorly beyond edge of ventral shield (Fig. 404); genital field of males with acetabula
borne in one or two rows on each side of gonopore (Fig. 404) ..............................................................................Chelohydracarus
89b.
Venter with coxal plates I not extending anteriorly beyond edge of ventral shield (Fig. 402); genital field of males with acetabula
borne in three or more rows on each side of gonopore (Figs. 400, 402) .......................................................................................90
90a(89b).
Genital field of males with all acetabula borne in gonopore (Fig. 402) ....................................................................Chelomideopsis
90b.
Genital field of males with acetabula borne both in gonopore and on acetabular plates flanking gonopore (Fig. 400)
...............................................................................................................................................................................Platyhydracarus
91a(84b).
Fourth pair of legs with tarsus indented or curved, but not greatly expanded, and bearing short peglike setae (Fig. 339)
..........................................................................................................................................Pionidae (in part) Foreliinae (in part) 92
91b.
Fourth pair of legs with tarsus not as described and illustrated above ..........................................................................................94
92a(91a).
Third pair of legs with tarsus and claws similar in shape to those of first two pairs of legs ..............................Pseudofeltria (males)
92b.
Third pair of legs with tarsus and claws modified, different from those on first two pairs of legs (Fig. 340).................................93
93a(92b).
Genital field with three pairs of acetabula (Fig. 336) ................................................................Pionacercus (males of some species)
93b.
Genital field with more than three pairs of acetabula (Fig. 338) .......................................................Forelia (males of some species)
94a(91b).
Fourth pair of legs with genu notched dorsally and bearing short peglike setae in notch (Fig. 343)..................................................
............................................................................................................................................Pionidae (in part) Pioninae (in part) 95
94b.
Fourth pair of legs with genu not notched as described and illustrated above ..............................................................................96
95a(94a).
Capitulum with anchoral process (Fig. 330, arrow A)........................................................................Piona (males of some species)
95b.
Capitulum lacking anchoral process (Fig. 341 and as shown in Fig. 342, arrow).............................................Nautarachna (males)
96a(94b).
Pedipalps with segments short and partially fused, femur with three ventral setae (Fig. 414, arrow)................................................
............................................................................................................................................Bogatiidae Horreolaninae Horreolanus
96b.
Pedipalps not as described and illustrated above ..........................................................................................................................97
97a(96b).
Genital field with three or four pairs of acetabula ........................................................................................................................98
97b.
Genital field with more than four pairs of acetabula ..................................................................................................................102
98a(97a).
Ventral shield completely surrounding gonopore; genital field with acetabula borne in gonopore (Fig. 407) or in pair of straight
or slightly curved rows on ventral shield flanking gonopore, when rows of acetabula slightly curved then genital field located well
anterior to posterior edge of idiosoma (Fig. 394) .........................................................................................................................99
98b.
Ventral shield not completely surrounding gonopore posteriorly (females of most genera) or completely surrounding gonopore
and with genital acetabula usually arranged in pronounced triangle or arc (males of all genera and females of one genus) (Fig.
368), when acetabula arranged in weak arc then genital field located at extreme posterior edge of ventral shield ......................112
99a(98a).
Dorsal shield with four pairs of glandularia including pair at anterior edge (Fig. 393, arrow); ventral shield with suture lines
between third and fourth coxal plates oriented at right angles to long axis of body where they meet midline (Fig. 394, arrow)
...............................................................................................................................Chappuisididae (in part) Morimotacarinae 100
16. Water Mites (Hydrachnida) and Other Arachnids
613
99b.
Dorsal shield with three pairs of glandularia (as in Fig. 411); ventral shield with suture lines between third and fourth coxal
plates oriented at oblique angle to long axis of body where they meet midline...........................................................................101
100a(99a).
Pedipalps with five segments (Fig. 391) ....................................................................................................................Morimotacarus
100b.
Pedipalps with four segments, including fused tibiotarsus (Fig. 392) .................................................................................Yachatsia
101a(99b).
Pedipalps with base of tarsus only about half as high as distal end of tibia, and inserted distidorsally on tibia (Fig. 417); ventral
shield with insertions of fourth pair of legs covered by projections of fourth coxal plates (Fig. 418, arrow) ....................................
...................................................................................................................................Acalyptonotidae (in part) Paenecalyptonotus
101b.
Pedipalps with base of tarsus approximately equal in height to distal end of tibia, and inserted distally on tibia (Fig. 405); ventral
shield with insertions of fourth pair of legs not covered by projections of fourth coxal plates (Fig. 407)..........................................
...............................................................................................................................................Mideopsidae Mideopsinae Mideopsis
102a(97b).
Dorsal shield bearing six pairs of glandularia, with three pairs grouped close together in an arc or triangle on each side (Fig. 322,
circled area) ................................................................................................Unionicolidae (in part) Pionatacinae (in part) Koenikea
102b.
Dorsal shield not as described and illustrated above...................................................................................................................103
103a(102b).
Gonopore located near middle of ventral shield (Fig. 406) .........................................................................................................104
103b.
Gonopore located near posterior end of ventral shield ...............................................................................................................105
104a(103a).
Ventral shield incorporating excretory pore (Fig. 406, arrow B) .......................................................................Laversiidae Laversia
104b.
Ventral shield not incorporating excretory pore...........Pionidae (in part) Pioninae (in part) Nautarachna (females of some species)
105a(103b).
Ventral shield with first coxal plates projecting beyond anterior edge of ventral shield (Figs. 377, 410, 412) .............................106
105b.
Ventral shield with first coxal plates not projecting beyond anterior edge of ventral shield (Fig. 357) ........................................123
106a(105a).
Pedipalps with femur and tibia bearing distinct ventral projections (Fig. 409, arrows); third and fourth pair of legs with welldeveloped swimming setae (Fig. 408).................................................................................................Amoenacaridae Amoenacarus
106b.
Pedipalps with femur and tibia usually lacking ventral projections, when bearing projections then third and fourth pair of legs
lack swimming setae...................................................................................................................................................................107
107a(106b).
Dorsal shield bearing three pairs of glandularia peripherally (Fig. 411) ..................................Arenohydracaridae Arenohydracarus
107b.
Dorsal shield bearing one to several pairs of glandularia, with at least one pair located well medial to edge (Fig. 376)....................
........................................................................................................................................................Aturidae (in part) Aturinae 108
108a(107b).
Dorsal shield bearing only one pair of glandularia (Fig. 379); pedipalps with segments extremely long and slender (Fig. 378)
......................................................................................................................................................................................Bharatalbia
108b.
Dorsal shield bearing three or more pairs of glandularia (Fig. 371); pedipalps with segments relatively short and stocky (Fig. 373)
...................................................................................................................................................................................................109
109a(108b).
Ventral shield bearing pronounced projections associated with openings of insertions of fourth legs (Figs. 374, 375)................110
109b.
Ventral shield lacking projections associated with openings of insertions of fourth legs (Fig. 372, 377) .....................................111
110a(109a).
Ventral shield with fourth coxal plates bearing pair of conspicuous glandularia medially (Fig. 374, arrow) ....................Neoaturus
110b.
Ventral shield with fourth coxal plates lacking glandularia (Fig. 375)...........................................................................Kongsbergia
111a(109b).
Ventral shield with genital acetabula in more than one row and restricted to region immediately lateral to gonopore near posterior edge of idiosoma (Fig. 372).........................................................................................................................Phreatobrachypoda
111b.
Ventral shield with genital acetabula in single row and distributed along posterolateral edge of shield between gonopore and insertions of fourth legs (Fig. 377) ............................................................................................................................................Aturus
112a(98b).
Dorsal shield comprising large central plate completely surrounded by ring of small, paired platelets (Fig. 347) .............................
....................................................................................................................................................Lethaxonidae (in part) Lethaxona
112b.
Dorsal shield comprising single large plate (although smaller posterior platelet bearing excretory pore may be present) (Fig. 365)
...................................................................................................................................Aturidae (in part) Axonopsinae (in part) 113
113a(112b).
Ventral shield with posterior edges of fourth coxal plates clearly indicated by suture lines that are indented posteromedially to
accommodate posterior coxoglandularia (Fig. 348, arrow) ....................................................................................................Ljania
113b.
Ventral shield with posterior edges of fourth coxal plates usually faintly indicated or obliterated by fusion with shield (Fig. 368)
and never indented posteromedially to accommodate posterior coxoglandularia ......................................................................114
614
Ian M. Smith, David R. Cook, Bruce P. Smith
114a(113b).
Fourth coxal plates with large projections associated with insertions of fourth pair of legs (Fig. 350, arrow A; Fig. 354, arrow)
...................................................................................................................................................................................................115
114b.
Fourth coxal plates with small projections or none associated with insertions of fourth pair of legs (Fig. 369) ..........................116
115a(114a).
Fourth coxal plates with pair of closely spaced glandularia in females (Fig. 349, arrow) and with looped suture lines extending
anteriorly in males (Fig. 350, arrow B) ...........................................................................................................................Stygalbiella
115b.
Fourth coxal plates with glandularia widely spaced in females (Fig. 351) and without looped suture lines in males (Fig. 352)
.....................................................................................................................................................................................Axonopsella
116a(114b).
Ventral shield lacking ridges in region anterolateral to insertions of fourth pair of legs (Figs. 355, 356) ....................................117
116b.
Ventral shield bearing well-developed ridges extending anterolaterally from region of insertion of fourth pair of legs (Fig. 368,
arrow) ........................................................................................................................................................................................118
117a(116a).
Ventral shield with anterior edges of first coxal plates extending to or beyond anterior edge of ventral shield (Fig. 355); genital
field with three pairs of acetabula (Fig. 355) ....................................................................................................................Albaxona
117b.
Ventral shield with anterior edges of first coxal plates not extending to anterior edge of ventral shield (Fig. 356); genital field with
four pairs of acetabula (Fig. 356) ........................................................................................................................................Javalbia
118a(116b).
Pedipalps with femur bearing spinelike projection in proximal three-quarters of ventral surface (Fig. 370); genital field of females
separated from ventral shield (Fig. 358) .....................................................................................................................................119
118b.
Pedipalps with femur lacking spinelike projection ventrally, although rounded projection may be present at distiventral extremity
(Fig. 366); genital field of females separated from or fused with ventral shield...........................................................................121
119a(118a).
Genital field bearing four pairs of acetabula (Fig. 369); fourth pair of legs of males not highly modified ................Neobrachypoda
119b.
Genital field bearing three pairs of acetabula (Fig. 358); fourth pair of legs of males highly modified .......................................120
120a(119b).
Fourth coxal plates bearing ridges extending posteriorly from region of insertions of fourth pair of legs (Fig. 358); fourth pair of
legs of males with tarsus expanded distally and flattened (Fig. 362) ..............................................................................Estellacarus
120b.
Fourth coxal plates lacking ridges extending posteriorly from region of insertions of fourth pair of legs (Fig. 359); fourth pair of
legs of males with tibia expanded and curved and tarsus slightly bowed and with enlarged claws (Fig. 363) ................Brachypoda
121a(118b).
Fourth coxal plates of males bearing prominent ridges extending posteriorly from region of insertions of fourth pair of legs (Fig.
364, arrow); fourth pair of legs of males with tibia and tarsus thicker and stockier than those of females (Fig. 367); genital field
of females separated from ventral shield ....................................................................................................................Woolastookia
121b.
Fourth coxal plates of males lacking prominent ridges extending posteriorly from region of insertions of fourth pair of legs (as in
Fig. 360), although short, faint ridges may be evident in some species (Fig. 368); fourth pair of legs of males similar to those of
females, either flattened or unmodified; genital field of females fused with ventral shield (Fig. 360) ...........................................122
122a(121b).
Fourth pair of legs with segments flattened and with telofemur relatively short and greatly expanded distally (Fig. 361).................
..................................................................................................................................................................................Erebaxonopsis
122b.
Fourth pair of legs with segments nearly cylindrical and with telofemur similar to other segments .................................Axonopsis
123a(105b).
Fourth coxal plates usually with projections partially covering insertions of fourth pair of legs, when projections absent then genital field with acetabula arranged in single rows extending laterally from gonopore in ventral shield (Fig. 346) ........................124
123b.
Fourth coxal plates lacking projections associated with insertions of fourth pair of legs and genital field with acetabula arranged
in several rows on acetabular plates (Fig. 357) ..............................................................................Aturidae (in part) Albiinae Albia
124a(123a).
Dorsal shield entire (Fig. 353); fourth coxal plates with projections associated with insertions of fourth pair of legs (Fig. 354, arrow).............................................................................................................Aturidae (in part) Axonopsinae (in part) Submiraxona
124b.
Dorsal shield comprising large central plate completely surrounded by ring of small, paired platelets (Fig. 345); fourth coxal
plates lacking projections associated with insertions of fourth pair of legs (Fig. 346)...............Lethaxonidae (in part) Lethaxonella
125a(71b).
Fourth coxal plates bearing glandularia (Fig. 319, arrow)......................................................................Hygrobatidae (in part) 126
125b.
Fourth coxal plates lacking glandularia......................................................................................................................................129
126a(125a).
Gnathosoma with capitulum broadly fused with first coxal plates (Fig. 315) .............................................................................127
126b.
Gnathosoma with capitulum separate from first coxal plates (Fig. 320) .....................................................................................128
127a(126a).
First pair of legs with tarsus straight (Fig. 317) ..............................................................................................................Hygrobates
127b.
First pair of legs with tarsus bowed (Fig. 318) .................................................................................................................Mesobates
128a(126b).
First coxal plates fused medially (Fig. 319); first pair of legs with distal segments modified as in Fig. 321........................Atractides
16. Water Mites (Hydrachnida) and Other Arachnids
615
128b.
First coxal plates separate medially (Fig. 320); first pair of legs with distal segments not modified ...............................Corticacarus
129a(125b).
Venter with apodemes of anterior coxal groups extending to approximately middle of fourth coxal plates (Fig. 323, arrow)
.................................................................................................................Unionicolidae (in part) Pionatacinae (in part) Neumania
129b.
Venter with apodemes of anterior coxal groups not extending to middle of fourth coxal plates .................................................130
130a(129b).
Genital field surrounded by well developed ventral shield (Fig. 418) .................................Acalyptonotidae (in part) Acalyptonotus
130b.
Genital field not surrounded by ventral shield ............................................................................................................................131
131a(130b).
Fourth coxal plates with medial edges reduced to angles (Fig. 338, arrow) ................................................................................132
131b.
Fourth coxal plates with medial edges not reduced to angles (Fig. 344, arrow) ..........................................................................135
132a(131a).
First pair of legs with tarsal claw socket large, occupying much more than one half dorsal surface of segment (Fig. 326, arrow);
genital field with three or four pairs of acetabula; fourth pair of legs with tarsus unmodified in males...............Wettinidae Wettina
132b.
First pair of legs with tarsal claw socket smaller, occupying one half or less of dorsal surface of segment; genital field with three
or more pairs of acetabula; fourth pair of legs with tarsus indented or bowed and bearing short peglike setae in males (Fig. 339)
........................................................................................................................................Pionidae (in part) Foreliinae (in part) 133
133a(132b).
Genital field with three pairs of acetabula (Fig. 336) .....................................................................................................Pionacercus
133b.
Genital field with more than three pairs of acetabula (Figs. 337, 338) .......................................................................................134
134a(133b).
Legs with swimming setae on distal segments ................................................Forelia (females of all species, males of some species)
134b.
Legs without swimming setae on distal segments ..........................................................................................Pseudofeltria (females)
135a(131b).
Capitulum without anchoral process (Fig. 342, arrow); symbionts of clams.................Pionidae (in part) Najadicolinae Najadicola
135b.
Capitulum with anchoral process (Fig. 330, arrow A); symbionts of clams or free living ...........................................................136
136a(135b).
Pedipalps with segments relatively long and slender (Fig. 329); genital field with three pairs of acetabula; males with petiole posterior to genital field (Fig. 327); and with genu of third pair of legs modified (Fig. 328)...................................................................
......................................................................................................................Pionidae (in part) Hydrochoreutinae Hydrochoreutes
136b.
Pedipalps with segments relatively short and stocky; genital field with three to many pairs of acetabula; males lacking petiole
posterior to genital field and with genu of third pair of legs unmodified.....................................................................................137
137a(136b).
Venter with suture lines between third and fourth coxal plates extending only a short distance toward midline (Fig. 324, arrow
A); with a pair of glandularia present at, and usually fused with, posteromedial angles of fourth coxal plates (Fig. 324, arrow B);
first pair of legs with long movable setae in free-living species (Fig. 325); symbionts of clams or sponges, or free living .................
............................................................................................................................Unionicolidae (in part) Unionicolinae Unionicola
137b.
Venter with suture lines between third and fourth coxal plates extending to (or nearly to) medial margins of coxal plates (as in
Fig. 330, usually longer than shown in Fig. 344); without glandularia at posteromedial angles of fourth coxal plates; first pair of
legs lacking long, movable setae; free living................................................................................................................................138
138a(137b).
Genital field usually with three, but rarely up to six, pairs of acetabula ........................................Pionidae (in part) Tiphyinae 139
138b.
Genital field with seven or more pairs of acetabula ...................................................................................................................142
139a(138a).
Fourth pair of legs unmodified ............................................................................Tiphyinae (females) Neotiphys, Pionopsis, Tiphys
139b.
Fourth pair of legs with segments modified....................................................................................................Tiphyinae (males) 140
140a(139b).
Fourth pair of legs with tibia bearing long, thick, curved seta dorsally (Fig. 333, arrow) and with genu only slightly expanded
.........................................................................................................................................................................................Neotiphys
140b.
Fourth pair of legs with tibia lacking long, thick, curved seta dorsally and with genu slightly or greatly expanded....................141
141a(140b).
Fourth pair of legs with genu greatly expanded and bearing numerous long setae with enlarged bases (Fig. 335)..................Tiphys
141b.
Fourth pair of legs with genu slightly expanded and bearing short setae with normal bases (Fig. 334)..............................Pionopsis
142a(138b).
Pedipalps with genu bearing long seta medially (Fig. 331, arrow); venter with second and third coxal plates each with pair of
thickened setae (Fig. 330, arrows B); legs of males unmodified.....................................Pionidae (in part) Huitfeldtiinae Huitfeldtia
142b.
Pedipalps with genu lacking long seta medially; venter with second and third coxal plates lacking thick setae (Fig. 344); fourth
pair of legs in males with genu notched and with peglike setae associated with notch (Fig. 343)......................................................
..........................................................................................................................................Pionidae (in part) Pioninae (in part) 143
143a(142b).
Capitulum with anchoral process (as in Fig. 330, arrow A) ..................................................Piona (females, males of most species)
143b.
Capitulum without anchoral process (Fig. 341 and as in Fig. 342, arrow) ...........................Nautarachna (females of some species)
616
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 200 Limnesia sp. (Limnesiinae), adult female, ventral view.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 201, 202 Stygothrombium sp. (Stygothrombidiidae) adult. Fig. 201, claws and empodium of leg I;
Fig. 202, pedipalp. FIGURES 203, 205–206 Hydrovolzia marshallae Cook (Hydrovolziinae), adult. Fig.
203, glandularium; Fig. 205, coxal plate III ; Fig. 206, dorsum of idiosoma. FIGURE 204 Acherontacarus sp.
(Acherontacarinae), adult, dorsum of idiosoma. FIGURES 207–209 Hydrachna sp. (Hydrachnidae), adult.
Fig. 207, dorsal view of idiosoma; Fig. 208, capitulum and chelicera; Fig. 209, pedipalp.
617
618
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 210 Eylais sp. (Eylaidae), adult, eye plate. FIGURES 211, 216 Limnochares sp. (Limnocharinae),
adult. Fig. 211, eye plate; Fig. 216, pedipalp. FIGURE 212 Piersigia limnophila Protz (Protziidae), adult,
eye plate. FIGURES 213–214 Apheviderulix santana Gerecke, Smith and Cook. (Apheviderulicidae),
deutonymph. Fig. 213 dorsum of idiosoma; Fig. 214, venter of idiosoma and gnathosoma. FIGURE 215
Rhyncholimnochares sp. (Rhyncholimnocharinae), adult, pedipalp. FIGURE 217 Neolimnochares sp. (Limnocharinae), adult, pedipalp. FIGURE 218 Clathrosperchon sp. (Clathrosperchontinae), adult, dorsum of
idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 219 Hydryphantes sp. (Hydryphantinae), adult, eye plate. FIGURE 220 Hydryphantes ruber (de
Geer) (Hydryphantinae), adult, dorsum of idiosoma. FIGURE 221 Hydrodroma sp. (Hydrodromidae), adult,
pedipalp. FIGURES 222, 228 Protzia sp. (Protziinae), adults. Fig. 222, claws of leg I; FIG. 228, genital
field. FIGURE 223 Cowichania interstitialis Smith (Cowichaniinae), adult female, venter of idiosoma.
FIGURE 224 Partnunia steinmanni Walter (Protziinae), adult, dorsum of idiosoma. FIGURES 225, 229
Notopanisus sp. (Thyadinae), adult. Fig. 225, medial eye; Fig. 229, genital field. FIGURES 226–227
Wandesia sp. (Wandesiinae), adult; Fig. 226, venter of idiosoma and gnathosoma; Fig. 227 pedipalp.
619
620
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 230, 231
Albertathyas montana Smith and Cook (Thyadinae), adult. Fig. 230, Dorsum of
idiosoma; Fig. 231, genital field. FIGURE 232 Cyclothyas siskiyouensis Smith (Cyclothyadinae), adult,
dorsum of idiosoma. FIGURE 233 Siskiyouthyas rivularis Smith and Cook (Thyadinae), adult, dorsum of
idiosoma. FIGURE 234 Trichothyas muscicola Mitchell (Thyadinae), adult, dorsum of idiosoma. FIGURE
235 Euthyas truncata (Neumann) (Thyadinae), adult, dorsum of idiosoma. FIGURE 236, 237 Euthyas
mitchelli Cook (Thyadinae), adult. Fig. 236, Genital field; Fig. 237, venter of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 238, 239 Thyopsis cancellata (Protz) (Thyadinae), adult. Fig. 238, Dorsal shield; Fig. 239, genital
field. FIGURE 240 Thyopsella occidentalis Cook (Thyadinae), adult, dorsal shield. FIGURES 241, 246
Thyopsoides plana (Habeeb) (Thyadinae), adult. Fig. 241, Dorsum of idiosoma; Fig. 246, genital field.
FIGURE 244 Thyopsella dictyophora Cook (Thyadinae), adult, genital field. FIGURE 242 Panisus
condensatus Habeeb (Thyadinae), adult, dorsum of idiosoma. FIGURE 243 Columbiathyas crenicola Smith
and Cook (Thyadinae), adult, dorsum of idiosoma. FIGURES 245, 247 Panisopsis setipes (Viets)
(Thyadinae), adult. Fig. 245, dorsum of idiosoma; Fig. 247, genital field.
621
622
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 248, 249
Parathyasella heatherensis Smith and Cook (Thyadinae), adult. Fig. 248, Dorsum of
idiosoma; Fig. 249, genital field. FIGURE 250 Amerothyasella appalachiana Smith and Cook (Thyadinae),
adult, dorsum of idiosoma. FIGURE 251 Thyasella sp. (Thyadinae), adult, dorsum of idiosoma. FIGURE
252 Thyasides sp. (Thyadinae), adult, dorsum of idiosoma. FIGURE 253 Thyas rivalis Koenike
(Thyadinae), adult, genital field. FIGURE 254 Tartarothyas sp. (Tartarothyadinae), adult, genital field.
FIGURE 255 Thermacarus nevadensis Marshall (Thermacaridae), adult, pedipalp. FIGURE 256 Thyas
stolli Koenike (Thyadinae),adult, dorsum of idiosoma. FIGURE 257 Zschokkea sp. (Thyadinae), adult,
genital field.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 258 – 260 Sperchon spp. (Sperchontidae), adults. Fig. 258, Dorsum of idiosoma; Fig. 259, venter
of idiosoma; Fig. 260, pedipalp. FIGURES 261, 262 Sperchonopsis nova Prasad and Cook (Sperchontidae),
adult. Fig. 261, Dorsum of idiosoma; Fig. 262, pedipalp. FIGURES 263, 264 Rutripalpus spp. (Rutripalpidae),
adult females. Fig. 263, Pedipalp; Fig. 264, venter of idiosoma. FIGURE 265 Teutonia sp. (Teutoniidae), adult,
venter of idiosoma.
623
624
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 266, 268 Utaxatax ovalis Cook (Anisitsiellinae), adult. Fig. 266, Tarsus of leg IV; Fig. 268, male,
ventral shield. FIGURE 267 Mamersellides sp. (Anisitsiellinae), adult, leg. IV. FIGURE 269 Bandakiopsis
fonticola Smith (Anisitsiellinae), adult, ventral shield. FIGURE 270 Cookacarus columbiensis Barr
(Anisitsiellinae), adult male, ventral shield. FIGURE 271 Bandakia phreatica Cook (Anisitsiellinae),
adult male, ventral shield. FIGURE 272 Oregonacarus rivulicolus Smith (Anisitsiellinae), adult male,
ventral shield.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 273, 274 Testudacarus americanus Marshall (Testudacarinae), adults. Fig. 273, Dorsal shield; Fig.
274, pedipalp. FIGURES 275 – 277 Neoatractides sp. (Neoatractidinae), adults. Fig. 275, Pedipalp; Fig. 276,
dorsal shield; Fig. 277, ventral shield. FIGURES 278, 279, 281 Torrenticola sp. (Torrenticolinae), adults.
Fig. 278, Dorsal shield; Fig. 279, ventral shield; Fig. 281, gnathosoma. FIGURE 280 Pseudotorrenticola
chiricahua Smith (Torrenticolinae), adult, gnathosoma. FIGURE 282 Monoatractides sp. (Torrenticolinae),
adult, gnathosoma.
625
626
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 283, 284 Lebertia sp. (Lebertiidae), adults. Fig. 283, Venter of idiosoma; Fig. 284, pedipalp.
FIGURE 285 Estelloxus montanus Habeeb (Lebertiidae) adult, venter of idiosoma. FIGURES 286, 287
Scutolebertia trinitensis Smith (Lebertiidae), adults. Fig. 286, Dorsum of idiosoma; Fig. 287, pedipalp.
FIGURES 288–290 Oxus spp. (Oxidae), adults. Fig. 288, Dorsum of idiosoma; Fig. 289, venter of
idiosoma; Fig. 290, tibia and tarsus of leg IV. FIGURE 291 Frontipoda americana Marshall (Oxidae), adult
male, dorsum of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 292 Omartacarus sp. (Omartacaridae), adult female, venter of idiosoma. FIGURE 293 Diamphidaxona sp. (Hygrobatidae), adult male, venter of idiosoma. FIGURE 294 Neomamersa chihuahua Smith and
Cook (Neomamersinae), adult female, dorsum of idiosoma. FIGURES 295, 299, 301 Neomamersa spp.
(Neomamersinae), adult males. Fig. 295, Pedipalp; Figs, 299, 301, genital fields. FIGURE 296 Kawamuracarus sp. (Kawamuracarinae), adult, pedipalp. FIGURES 297, 303 Arizonacarus chiricahuensis Smith and
Cook (Neomamersinae), adult females. Fig. 297, Pedipalp; Fig. 303, venter of idiosoma. FIGURES 298, 300,
304 Meramecia occidentalis Smith and Cook (Neomamersinae), adult females. Fig. 298, Pedipalp; Fig. 300,
genital field; Fig. 304, venter of idiosoma. FIGURE 302 Neomamersa californica Smith and Cook
(Neomamersinae), adult female, venter of idiosoma.
627
628
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 305 Centrolimnesia bondi Lundblad (Limnesiinae), adult male, genital field. FIGURES 306,
309 – 311 Tyrrellia spp. (Tyrelliinae), adults. Fig. 306, Pedipalp; Fig. 309, male, venter of idiosoma; Fig. 310,
male, dorsal shield; Fig. 311, male, dorsum of idiosoma. FIGURE 307 Protolimnesia sp. (Protolimnesiinae),
adult, dorsum of idiosoma. FIGURE 308 Neotyrrellia anitahoffmannae Smith and Cook (Tyrrellinae), adult
female, venter of idiosoma. FIGURE 312 Feltria exilis Cook (Feltriidae), adult female, venter of idiosoma.
FIGURES 313, 314 Frontipodopsis sp. (Frontipodopsidae), adult. Fig. 313, Leg IV; Fig. 314, lateral view
of idiosoma.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 315 – 317
Hygrobates spp. (Hygrobatidae), adults. Fig. 315 male, venter of idiosoma; Fig. 316 ,
Chelicera; Fig. 317, tibia and tarsus of leg I. FIGURE 318 Mesobates sp. (Hygrobatidae), adult, tibia and
tarsus of leg I. FIGURES 319, 321 Atractides sp. (Hygrobatidae), adults. Fig. 319, Male, venter of idiosoma;
Fig. 321, tibia and tarsus of leg I. FIGURE 320 Corticacarus sp. (Hygrobatidae), adult female, venter of
idiosoma. FIGURE 322 Koenikea sp. (Pionatacinae), adult dorsal shield. FIGURE 323 Neumania sp.
(Pionatacinae), adult male, venter of idiosoma. FIGURES 324, 325 Unionicola sp. (Unionicolinae), adults.
Fig. 324, female, venter of idiosoma; Fig. 325, leg I.
629
630
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 326
Wettina octopora Cook (Wettinidae), adult, leg I. FIGURES 327 – 329 Hydrochoreutes
intermedius Cook (Hydrochoreutinae), adults. Fig. 327, Male, genital field; Fig. 328, male, genu of leg III;
Fig. 329, pedipalp. FIGURES 330, 331 Huitfeldtia rectipes (Huitfeldtiinae), adults. Fig. 330, female, venter
of idiosoma; Fig. 331, pedipalp. FIGURES 332, 335 Tiphys spp. (Tiphyinae), adults. Fig. 332, pedipalp; Fig.
335, male, leg IV. FIGURE 333 Neotiphys pionoidellus (Habeeb) (Tiphyinae), adult male, tibia of leg IV.
FIGURE 334 Pionopsis paludis Habeeb (Tiphyinae), adult male, leg IV.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 336 Pionacercus sp. (Foreliinae), adult male, venter of idiosoma. FIGURE 337 Pseudofeltria
multipora Cook (Foreliinae), adult male, ventral shield. FIGURES 338 – 340 Forelia floridensis Cook
(Foreliinae), adult male. Fig. 338, Venter of idiosoma; Fig. 339, distal segments of leg IV; Fig. 340, tibia and
tarsus of leg III. FIGURE 341 Nautarachna queticoensis Smith (Pioninae), adult female, gnathosoma.
FIGURE 342 Najadicola ingens Koenike (Najadicolinae), adult male, venter of idiosoma. FIGURES 343,
344 Piona spp. (Pioninae), adult males. Fig. 343, Genu of leg IV; Fig. 344, venter of idiosoma.
631
632
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 345, 346 Lethaxonella parva Cook (Lethaxonidae), adult female. Fig. 345, Dorsal plate; Fig. 346,
ventral plate. FIGURE 347 Lethaxona sp. (Lethaxonidae), adult, dorsal shield. FIGURE 348 Ljania
michiganensis Cook (Axonopsinae), adult male, ventral shield. FIGURES 349, 350 Stygalbiella yavapai
Smith and Cook (Axonopsinae), adults. Fig. 349, Female, ventral shield; Fig. 350, male, ventral shield.
FIGURES 351, 352 Axonopsella bakeri Smith and Cook (Axonopsinae), adults. Fig. 351, female, ventral
shield; Fig. 352, male, ventral shield.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 353, 354 Submiraxona gilana (Habeeb) (Axonopsinae), adults. Fig. 353, Dorsal shield; Fig. 354,
male, ventral shield. FIGURE 355 Albaxona nearctica Cook (Axonopsinae), adult male, ventral shield.
FIGURE 356 Javalbia sp. (Axonopsinae), adult female, ventral shield. FIGURE 357 Albia sp. (Albiinae),
adult male, ventral shield. FIGURE 358 Estellacarus unguitarsus (Habeeb) (Axonopsinae), adult female,
ventral shield. FIGURE 359 Brachypoda oakcreekensis Habeeb (Axonopsinae), adult female, ventral shield.
FIGURES 360, 361 Erebaxonopsis nearctica Cook (Axonopsinae), adults. Fig. 360, female,ventral shield;
Fig. 361, male, leg IV.
633
634
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 362 Estellacarus unguitarsus (Habeeb) (Axonopsinae), adult male, distal segments of leg IV.
FIGURE 363 Brachypoda oakcreekensis Habeeb (Axonopsinae), adult male, distal segments of leg IV.
FIGURES 364 – 367 Woolastookia pilositarsa (Habeeb) (Axonopsinae), adults. Fig. 364, Male, venter of
idiosoma; Fig. 365, female, pedipalp; Fig. 366, female, dorsal shield; Fig. 367, male, distal segments of leg IV.
FIGURE 368 Axonopsis beltista Cook (Axonopsinae), adult male, ventral shield. FIGURES 369, 370
Neobrachypoda ekmani (Walter) (Axonopsinae), adults. Fig. 369, Female, venter of idiosoma; Fig. 370,
male, pedipalp.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 371, 372 Phreatobrachypoda multipora Cook (Aturinae), adult male. Fig. 371, Dorsum of
idiosoma; Fig. 372, venter of idiosoma. FIGURE 373 Phreatobrachypoda oregonensis Smith (Aturinae),
adult male, pedipalp. FIGURE 374 Neoaturus sp. (Aturinae), adult female, venter of idiosoma. FIGURES
375, 376 Kongsbergia spp. (Aturinae), adult males. Fig. 375, Venter of idiosoma and fourth leg; Fig. 376,
dorsal shield. FIGURE 377 Aturus sp. (Aturinae), adult female, venter of idiosoma. FIGURES 378, 379
Bharatalbia spp. (Aturinae), adults. Fig. 378, B. cooki Smith, female, dorsum of idiosoma; Fig. 379, B. surensis
Smith, male, pedipalp.
635
636
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURE 380
Cyclomomonia andrewi Smith (Cyclomomoniinae), adult, dorsal shield. FIGURE 381
Stygomomonia riparia Habeeb (Stygomomoniinae), adult, distal segments of leg I. FIGURE 382 Momonia
projecta Cook (Momoniinae), adult female, ventral shield. FIGURE 383 Stygomomonia mitchelli Smith
(Stygomomoniinae), adult male, ventral shield. FIGURE 384 Midea expansa Marshall (Mideidae), adult
female, ventral shield. FIGURE 385 Nudomideopsis magnecetabula (Smith) (Nudomideopsidae), adult male,
ventral shield. FIGURE 386 Paramideopsis susanae Smith (Nudomideopsidae), adult male, ventral shield.
FIGURE 387 Neomideopsis suislawensis Smith (Nudomideopsidae), adult male, ventral shield.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURE 388
Chappuisides eremitus Cook (Chappuisidinae), adult male, ventral shield. FIGURE 389
Uchidastygacarus ovalis Cook (Uchidastygacarinae), adult female, ventral shield and legs. FIGURE 390
Uchidastygacarus acadiensis Smith (Uchidastygacarinae), adult, pedipalp. FIGURE 391 Morimotacarus
nearcticus Smith (Morimotacarinae), adult, pedipalp. FIGURES 392 – 394 Yachatsia mideopsoides Cook
(Morimotacarinae), adults. Fig. 392, Pedipalp; Fig. 393, dorsal shield; Fig. 394, female, ventral shield.
FIGURES 395, 397 Volsellacarus sabulonus Cook (Neoacaridae), adults. Fig. 395, female, ventral shield;
Fig. 397, gnathosoma. FIGURE 396 Neoacarus minimus Cook (Neoacaridae), adult, pedipalp.
637
638
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 398, 399
Stygameracarus cooki Smith (Stygameracarinae), adults. Fig. 398, Male, ventral shield;
Fig. 399, pedipalp. FIGURE 400 Platyhydracarus juliani Smith (Athienemanniinae), adult male, genital
field. FIGURES 401, 402 Chelomideopsis besselingi (Cook) (Athienemanniinae), adults. Fig. 401,
Gnathosoma; Fig. 402, adult male, ventral shield. FIGURES 403, 404 Chelohydracarus navarrensis
Smith (Athienemanniinae), adult male. Fig. 403, Genital field; Fig. 404, ventral shield. FIGURES 405, 407
Mideopsis sp. (Mideopsinae), adults. Fig. 405, Pedipalp; Fig. 407, female, ventral shield. FIGURE 406
Laversia berulophila Cook (Laversiidae), adult female, ventral shield.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 408 – 410 Amoenacarus dixiensis Smith and Cook (Amoenacaridae), adults. Fig. 408, Male, leg IV;
Fig. 409, pedipalp; Fig. 410, male, ventral shield. FIGURES 411, 412 Arenohydracarus minimus Cook
(Arenohydracaridae), adults. Fig. 411, dorsal shield; Fig. 412, female, ventral shield. FIGURE 413 Geayia sp.
(Krendowskiidae), adult female, ventral shield. FIGURE 414 Horreolanus orphanus Mitchell (Horreolaninae),
adult, pedipalp. FIGURES 415, 416 Arrenurus sp. (Arrenurinae), adults. Fig. 415, Female, ventral shield.
Fig. 416, pedipalp. FIGURES 417, 418 Acalyptonotus neoviolaceus Smith (Acalyptonotidae), adults.
Fig. 417, Pedipalp; Fig. 418, female, ventral shield.
639
640
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 419 – 422 Scanning electron micrographs of adult water mites. Fig. 419 Arrenurus fissicornis
Marshall (Arrenuruidae), male; Fig. 420 Arrenurus (Megaluracarus) pseudocylindratus Marshall (Arrenuridae),
male; Fig. 421 Kongsbergia sp. (Aturidae), female; Fig. 422 Aturus sp. (Aturidae), male.
16. Water Mites (Hydrachnida) and Other Arachnids
FIGURES 423 – 426 Scanning electron micrographs of adult water mites. FIGURE. 423 Koenikea wolcotti
Viets (Unionicolidae), female; FIGURE 424 Uchidastygacarus acadiensis Smith (Chappuisididae), female;
FIGURE 425 Paramideopsis susanae Smith (Nudomideopsidae), female; FIGURE 426 Volsellacarus
sabulonus Cook (Neoacaridae), female.
641
642
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 427 – 430 Photographs of parasitic larval water mites attached to host insects. FIGURE 427
Larvae of Eylais sp. attached to the subelytral abdominal integument of adult waterboatmen (Hemiptera:
Corrixidae) illustrating larval size before and after engorgement; FIGURE 428 larvae of Hydrachna sp.
(Hydrachnidae) attached to the thorax and elytra of a giant water bug (Hemiptera:Belostomatidae); Fig. 429
larvae of Arrenurus sp. (Arrenuridae) attached to the abdominal segments of a damselfly (Odonata:Zygoptera);
FIGURES 430 larvae of Feltria sp. (Feltriidae) and Aturus sp. (Aturidae) attached to the abdominal segments
of a chironomid midge (Diptera:Chironomidae).
TABLE I
Summary of Some Important Zoogeographic and Ecological Characteristics of North American Water Mite Genera
Ecology
Zoogeography
Taxa
Stygothrombidioidea
Stygothrombidiidae
Stygothrombium
Hydrovolzioidea
Hydrovolziidae
Hydrovolzia
Acherontacaridae
Acherontacarus
Hydrachnoidea
Hydrachnidae
Hydrachna
Eylaoidea
Limnocharidae
Limnocharinae
Limnochares
Neolimnochares
Rhyncholimnocharinae
Rhyncholimnochares
Eylaidae
Eylais
Piersigiidae
Piersigia
Apheviderulicidae
Apheviderulix
Hydryphantoidea
Hydryphantidae
Hydryphantinae
Hydryphantes
643
Thyadinae
Albertathyas
Amerothyasella
Columbiathyas
Larvae
Deutonymphs and Adults
Adults
Origin
North American
distribution
Type
Order of
hosts
Attachment site
Habitat type
Mode of
locomotion
Prey
Spermatophore
transfer mode
Laurasian
Temperate, boreal
Aquatic
Plecoptera
Thorax, abdomen
Interstitial
Crawling
Unknown
Unknown
Laurasian
Temperate, boreal
Terrestrial
Hemiptera,
Diptera
Thorax, abdomen
Springs, riffles
Crawling
Unknown
Unknown
Laurasian
Temperate
Aquatic
Unknown
Unknown
Interstitial
Crawling
Unknown
Unknown
Pangean
Throughout
Aquatic
Coleoptera,
Hermiptera
Thorax, abdomen
Lakes, temporary
pools
Swimming
Insect eggs
Dissociation
Pangean
Temperate, boreal
Terrestrial
Thorax, abdomen
Terrestrial
Thorax, abdomen
Crawling,
swimming
Crawling
UnKnown
Temperate
Riffles, pools,
lakes
Pools
Diptera larvae
Pangean
Hemiptera,
Odonata
Hemiptera
Unknown
Unknown
Gondwanan
Temperate
Aquatic
Coleoptera
Abdomen
Riffles, interstitial
Crawling
Unknown
Unknown
Pangean
Throughout
Terrestrial
Coleoptera,
Hemiptera
Thorax, abdomen
Pools, lakes,
temporary pools
Swimming
Ostracoda,
Cladocera
Dissociation,
association,
copulation
Laurasian
Temperate, boreal
Terrestrial
Coleoptera
Abdomen
Springs, interstitial,
temporary pools
Crawling
Ostracoda
Unknown
Laurasian
Temperate
Terrestrial
Coleoptera
Abdomen
Springs
Crawling
Unknown
Unknown
Pangean
Throughout
Terrestrial
Hemiptera,
Odonata,
Diptera
Thorax, abdomen
Lakes, temporary
pools
Swimming
Diptera eggs
Dissociation
North American
North American
North American
Temperate
Temperate
Temperate
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Riffles
Riffles
Springs
Crawling
Crawling
Crawling
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
(continues)
644
TABLE I
(Continued)
Ecology
Zoogeography
Larvae
Deutonymphs and Adults
Origin
North American
distribution
Type
Order of
hosts
Attachment site
Habitat type
Euthyas
Laurasian
Temperate
Terrestrial
Diptera
Thorax
Notopanisus
Panisus
Pangean
Laurasian
Temperate
Temperate, boreal
Unknown
Terrestrial
Unknown
Thorax
Panisopsis
Parathyasella
Siskiyouthyas
Thyas
Laurasian
Laurasian
North American
Laurasian
Temperate, boreal
Temperate, boreal
Temperate
Temperate, boreal
Terrestrial
Unknown
Unknown
Terrestrial
Thyasella
Thyasides
Thyopsella
Laurasian
Laurasian
North American
Temperate
Boreal
Temperate, boreal
Unknown
Terrestrial
Terrestrial
Thyopsis
Laurasian
Temperate
Terrestrial
Unknown
Diptera,
Hymenoptera
Diptera
Unknown
Unknown
Collembola,
Diptera,
Trichoptera
Unknown
Diptera
Diptera,
Trichoptera
Diptera
Riffles, temporary
pools
Springs, riffles
Springs, riffles
Thyopsoides
Trichothyas
North American
Laurasian
Temperate
Temperate
Unknown
Terrestrial
Unknown
Diptera
Laurasian
Boreal, arctic
Unknown
Unknown
Laurasian
Laurasian
Temperate
Temperate, boreal
Terrestrial
Terrestrial
Plecoptera
Diptera,
Trichoptera
Pangean
Temperate
Aquatic
Pangean
Temperate
Laurasian
Taxa
Zschokkea
Protziinae
Partnunia
Protzia
Wandesiinae
Wandesia
Tartarothyadinae
Tartarothyas
Cyclothyadinae
Cyclothyas
Cowichaniinae
Cowichania
Hydrodromidae
Hydrodroma
Rhynchohydracaridae
Clathrosperchoninae
Clathrosperchon
Adults
Mode of
locomotion
Prey
Spermatophore
transfer mode
Walking
Insect eggs
Unknown
Crawling
Crawling
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Thorax
Springs, riffles
Springs, riffles
Riffles
Springs, riffles,
temporary pools
Crawling
Crawling
Unknown
Walking
Unknown
Unknown
Unknown
Insect eggs
Unknown
Unknown
Unknown
Dissociation
Unknown
Thorax
Thorax, abdomen
Springs, interstitial
Temporary pools
Springs, riffles
Walking
Walking
Crawling
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Thorax
Springs, temporary
pools
Riffles
Springs
Crawling
Unknown
Unknown
Crawling
Walking
Unknown
Unknown
Unknown
Unknown
Temporary pools
Walking
Unknown
Unknown
Thorax, abdomen
Thorax, abdomen
Riffles
Riffles
Walking
Walking
Unknown
Unknown
Unknown
Unknown
Plecoptera
Thorax
Springs, interstitial
Crawling
Unknown
Unknown
Terrestrial
Unknown
Unknown
Springs, riffles
Walking,
crawling
Unknown
Unknown
Temperate (W)
Unknown
Unknown
Unknown
Springs, interstitial
Crawling
Unknown
Unknown
North American
Temperate (W)
Unknown
Unknown
Unknown
Interstitial
Crawling
Unknown
Unknown
Pangean
Throughout
Terrestrial
Diptera
Thorax
Riffles, pools, lakes
Swimming
Insect eggs
Dissociation
Gondwanan
Temperate
Unknown
Unknown
Unknown
Riffles, interstitial
Crawling
Unknown
Unknown
Unknown
Head, thorax,
abdomen
Unknown
Thermacaridae
Thermacarus
Pangean
Temperate
Terrestrial
Anura
(Amphibia)
Body dorsum
Hot springs
Crawling
Diptera larvae
Unknown
Lebertioidea
Sperchontidae
Sperchontinae
Sperchon
Laurasian
Throughout
Aquatic
Thorax, abdomen
Diptera larvae
Unknown
Temperate, boreal
Aquatic
Thorax
Springs, riffles,
pools, lakes
Riffles, pools, lakes
Crawling
Laurasian
Diptera,
Trichoptera
Diptera
Crawling
Unknown
Unknown
Laurasian
Boreal
Aquatic
Diptera
Abdomen
Springs, pools
Crawling
swimming
Diptera larvae
Unknown
Laurasian
Boreal (E)
Unknown
Unknown
Unknown
Springs, pools
Crawling
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Thorax
Crawling
Unknown
Unknown
Bandakiopsis
Cookacarus
Mamersellides
Oregonacarus
Utaxatax
Lebertiidae
Estelloxus
Lebertia
Laurasian
North American
Gondwanan
North American
Laurasian
Temperate
Temperate
Temperate (SE)
Temperate
Temperate
Aquatic
Unknown
Unknown
Unknown
Aquatic
Diptera
Unknown
Unknown
Unknown
Diptera
Thorax, abdomen
Unknown
Unknown
Unknown
Thorax, abdomen
Springs, riffles,
interstitial
Springs
Springs, interstitial
Pools
Springs
Springs, interstitial
Crawling
Crawling
Swimming
Crawling
Crawling
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
North American
Laurasian
Temperate (W)
Throughout
Aquatic
Aquatic
Diptera
Diptera
Thorax
Thorax
Unknown
Dissociation
North American
Temperate (SW)
Unknown
Unknown
Unknown
Walking
Walking,
swimming
Crawling
Unknown
Diptera larvae
Scutolebertia
Oxidae
Frontipoda
Oxus
Torrenticolidae
Neoatractidinae
Neoatractides
Testudacarinae
Testudacarus
Torrenticolinae
Monatractides
Pseudotorrenticola
Torrenticola
Springs, riffles
Springs, riffles,
pools, lakes
Springs
Unknown
Unknown
Pangean
Pangean
Temperate, boreal
Temperate, boreal
Aquatic
Aquatic
Diptera
Diptera
Thorax
Thorax, abdomen
Pools, lakes
Pools, lakes
Swimming
Swimming
Diptera larvae
Diptera larvae
Unknown
Unknown
Gondwanan
Temperate (SW)
Unknown
Unknown
Unknown
Riffles
Crawling
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Thorax
Riffles, interstitial
Crawling
Diptera larvae
Unknown
Laurasian
Laurasian
Laurasian
Temperate
Temperate
Temperate, boreal
Aquatic
Unknown
Aquatic
Diptera
Unknown
Diptera
Thorax
Unknown
Thorax
Riffles, pools, lakes
Interstitial
Riffles, interstitial,
pools, lakes
Crawling
Crawling
Crawling
Diptera larvae
Unknown
Diptera larvae
Unknown
Unknown
Unknown
Hygrobatoidea
Limnesiidae
Kawamuracarinae
Kawamuracarus
Laurasian
Temperate (SW)
Unknown
Unknown
Unknown
Interstitial
Crawling,
walking
Unknown
Unknown
Limnesiinae
Centrolimnesia
South American
Temperate
Unknown
Unknown
Unknown
Pools
Swimming
Unknown
Unknown
Sperchonopsis
Teutoniidae
Teutonia
Rutripalpidae
Rutripalpus
Anisitsiellidae
Anisitsiellinae
Bandakia
645
(continues)
646
TABLE I
(Continued)
Ecology
Zoogeography
Larvae
Deutonymphs and Adults
Adults
Origin
North American
distribution
Type
Order of
hosts
Attachment site
Habitat type
Mode of
locomotion
Pangean
Temperate, boreal
Aquatic
Diptera
Thorax
Riffles, interstitial,
pools, lakes
Crawling,
swimming
Copepoda,
cladocera,
insect eggs and
larvae
Unknown
Neomamersinae
Arizonacarus
North American
Temperate (SW)
Unknown
Unknown
Unknown
Interstitial
Unknown
Unknown
Meramecia
South American
Temperate
Unknown
Unknown
Unknown
Interstitial
Unknown
Unknown
Neomamersa
South American
Temperate
Unknown
Unknown
Unknown
Interstitial
Crawling,
walking
Crawling,
walking
Crawling,
walking
Unknown
Unknown
South American
Temperate (SW)
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
South American
South American
Temperate
Temperate, boreal
Unknown
Aquatic
Unknown
Diptera
Unknown
Abdomen
Riffles
Springs, pools, lakes
Crawling
Crawling
Unknown
Diptera eggs
and larvae
Unknown
Unknown
Gondwanan
Temperate (SW)
Unknown
Unknown
Unknown
Interstitial
Crawling
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Crawling,
walking
Temperate
Temperate
Boreal, temperate
Unknown
Unknown
Aquatic
Unknown
Unknown
Abdomen
Laurasian
Boreal
Unknown
Unknown
Unknown
Diptera,
Trichoptera
Unknown
Unknown
Crawling
Walking
Crawling,
swimming
Crawling
Diptera larvae,
Ostracoda,
Cladocera
Unknown
Unknown
Diptera larvae,
Ostracoda
Unknown
Unknown
Gondwanan
Gondwanan
Laurasian
Springs, riffles,
interstitial, pools,
lakes
Riffles
Interstitial
Springs, riffles,
pools, lakes
Riffles, pools
Unknown
Pangean
Temperate
Aquatic
Abdomen
Pools, lakes
Swimming
Diptera larvae
Unknown
Neumania
Pangean
Temperate, boreal
Aquatic
Diptera,
Trichoptera
Diptera
Abdomen
Pools, lakes
Swimming
Diptera larvae,
Copepoda
Dissociation,
association
Unionicolinae
Unionicola
Pangean
Temperate, boreal
Aquatic
Diptera,
Trichoptera
Abdomen, legs
Pools, lakes,
mollusc parasites
Crawling,
swimming
Copepoda,
Cladocera,
Diptera larvae,
mollusc tissue
Dissociation,
association,
copulation
Taxa
Limnesia
Protolimnesiinae
Protolimnesia
Tyrrelliinae
Neotyrrellia
Tyrrellia
Omartacaridae
Omartacarinae
Omartacarus
Hygrobatidae
Atractides
Corticacarus
Diamphidaxona
Hygrobates
Mesobates
Unionicolidae
Pionatacinae
Koenikea
Prey
Spermatophore
transfer mode
Unknown
Unknown
Dissociation
Feltriidae
Feltria
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Springs, riffles,
interstitial
Crawling
Diptera larvae
Copulation
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Pools, lakes
Swimming
Unknown
Dissociation
Laurasian
North American
Temperate (W)
Temperate (SW)
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Interstitial
Interstitial
Walking
Walking
Unknown
Unknown
Unknown
Unknown
Pangean
Temperate (W)
Unknown
Unknown
Unknown
Interstitial
Walking,
swimming
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Pools, lakes
Diptera larvae
Copulation
Laurasian
Boreal
Aquatic
Diptera
Abdomen
Springs, pools
Unknown
Copulation
Laurasian
Temperate
Aquatic
Diptera
Abdomen
Springs, interstitial
Crawling,
swimming
Crawling,
swimming
Crawling
Diptera larvae
Copulation
Laurasian
Boreal
Aquatic
Diptera
Abdomen
Lakes (profundal)
Swimming
Cladocera
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Pools, lakes
Swimming
Cladocera
Unknown
Laurasian
Temperate
Aquatic
Diptera
Abdomen
Lakes (mollusc
parasite)
Crawling
Mollusc tissue
Unknown
Laurasian
Throughout
Aquatic
Diptera
Abdomen
Copulation
Throughout
Aquatic
Diptera
Abdomen
Crawling,
swimming
Crawling,
swimming
Diptera larvae
Laurasian
Springs, riffles,
pools
Pools, lakes,
temporary pools
Copepoda,
Cladocera,
Diptera larvae
Copulation
North American
Laurasian
Boreal, temperate
Boreal
Aquatic
Aquatic
Diptera
Diptera
Abdomen
Abdomen
Pools
Pools, temporary
pools
Swimming
Swimming
Copulation
Copulation
Tiphys
Laurasian
Throughout
Aquatic
Diptera
Abdomen
Pools, lakes,
temporary pools
Swimming
Unknown
Ostracoda,
Cladocera,
Copepoda
Diptera larvae,
Ostracoda,
Cladocera,
Copepoda
Aturidae
Albiinae
Albia
Aturinae
Aturus
Laurasian
Temperate
Aquatic
Trichoptera
Abdomen
Pools, lakes
Swimming
Unknown
Unknown
Laurasian
Temperate
Aquatic
Diptera
Abdomen
Crawling
Diptera larvae
Copulation
Laurasian
Laurasian
South American
Temperate (W)
Temperate
Temperate
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Springs, riffles,
interstitial, pools
Interstitial
Riffles
Riffles
Walking
Crawling
Crawling
Unknown
Unknown
Unknown
Unknown
Copulation
Unknown
Wettinidae
Wettina
Lethaxonidae
Lethaxona
Lethaxonella
Frontipodopsidae
Frontipodopsis
Pionidae
Foreliinae
Forelia
Pionacercus
Pseudofeltria
Huitfeldtiinae
Huitfeldtia
Hydrochoreutinae
Hydrochoreutes
Najadicolinae
Najadicola
Pioninae
Nautarachna
Piona
Tiphyinae
Neotiphys
Pionopsis
647
Bharatalbia
Kongsbergia
Neoaturus
Copulation
(continues)
648
TABLE I
(Continued)
Ecology
Zoogeography
Larvae
Deutonymphs and Adults
Adults
Origin
North American
distribution
Type
Order of
hosts
Attachment site
Habitat type
Mode of
locomotion
Prey
Spermatophore
transfer mode
Laurasian
Temperate
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
Laurasian
Gondwanan
Laurasian
Temperate (E)
Temperate (E)
Temperate, boreal
Unknown
Unknown
Aquatic
Unknown
Unknown
Diptera
Unknown
Unknown
Abdomen
Unknown
Unknown
Unknown
Laurasian
Laurasian
North American
Laurasian
Laurasian
Laurasian
Gondwanan
Gondwanan
Temperate, boreal
Temperate
Boreal
Temperate
Temperate, boreal
Boreal
Temperate (W)
Temperate (W)
Aquatic
Unknown
Aquatic
Unknown
Aquatic
Aquatic
Unknown
Unknown
Diptera
Unknown
Diptera
Unknown
Diptera
Unknown
Unknown
Unknown
Abdomen
Unknown
Abdomen
Unknown
Abdomen
Unknown
Unknown
Unknown
Diptera larvae
Unknown
Diptera larvae
Unknown
Diptera larvae
Unknown
Unknown
Unknown
Copulation
Unknown
Copulation
Unknown
Unknown
Unknown
Unknown
Unknown
Woolastookia
Laurasian
Temperate, boreal
Aquatic
Diptera
Abdomen
Riffles, pools, lakes
Walking
Walking
Walking,
swimming
Swimming
Walking
Swimming
Walking
Walking
Walking
Walking
Walking,
swimming
Walking,
swimming
Unknown
Unknown
Diptera larvae
Brachypoda
Erebaxonopsis
Estellacarus
Javalbia
Ljania
Neobrachypoda
Stygalbiella
Submiraxona
Interstitial
Interstitial
Riffles, interstitial,
pools, lakes
Pools
Interstitial
Pools, lakes
Interstitial
Riffles
Springs, pools
Interstitial
Riffles, pools
Diptera larvae
Copulation
Laurasian
Throughout
Aquatic
Diptera
Abdomen
Pools, lakes
Swimming
Diptera larvae
Copulation
North American
Temperate (W)
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
Laurasian
Temperate
Aquatic
Trichoptera
Thorax, abdomen
Pools, lakes
Swimming
Diptera larvae
Unknown
Laurasian
Temperate
Aquatic
Trichoptera
Abdomen
Interstitial
Walking
Unknown
Unknown
North American
Pangean
Pangean
Temperate (W)
Temperate, boreal
Temperate, boreal
Unknown
Aquatic
Aquatic
Unknown
Diptera
Diptera
Unknown
Thorax
Thorax
Springs
Springs, interstitial
Springs, interstitial
Crawling
Crawling
Crawling
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Thorax
Springs, riffles,
interstitial, pools,
lakes
Crawling,
walking,
swimming
Diptera larvae
Unknown
Taxa
Phreatobrachypoda
Axonopsinae
Albaxona
Axonopsella
Axonopsis
Arrenuroidea
Mideidae
Midea
Momoniidae
Cyclomomoniinae
Cyclomomonia
Momoniinae
Momonia
Stygomomoniinae
Stygomomonia
Nudomideopsidae
Neomideopsis
Nudomideopsis
Paramideopsis
Mideopsidae
Mideopsinae
Mideopsis
Chappuisididae
Chappuisidinae
Chappuisides
Morimotacarinae
Morimotacarus
Yachatsia
Uchidastygacarinae
Uchidastygacarus
Neoacaridae
Neoacarus
Volsellacarus
Bogatiidae
Horreolaninae
Horreolanus
Acalyptonotidae
Paenecalyptonotus
Acalyptonotus
Athienemanniidae
Athienemanniinae
Chelohydracarus
Chelomideopsis
Platyhydracarus
Stygameracarinae
Stygameracarus
Amoenacaridae
Amoenacarus
Arenohydracaridae
Arenohydracarus
Laversiidae
Laversia
Krendowskiidae
Geayia
Krendowskia
Arrenuridae
Arrenurinae
Arrenurus
Laurasian
Temperate
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
Laurasian
Laurasian
Temperate (W)
Temperate (W)
Unknown
Unknown
Unknown
Unknown
Unknown
Unknown
Interstitial
Interstitial
Walking
Walking
Unknown
Unknown
Unknown
Unknown
Laurasian
Temperate
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
Laurasian
Temperate, boreal
Aquatic
Diptera
Thorax
Walking
Unknown
Unknown
North American
Temperate
Aquatic
Diptera
Thorax
Riffles, interstitial,
lakes
Riffles, interstitial
Walking
Unknown
Unknown
North American
Temperate
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
North American
Laurasian
Boreal
Boreal
Aquatic
Aquatic
Unknown
Diptera
Unknown
Thorax
Springs
Springs, pools, lakes
Crawling
Walking
Unknown
Unknown
Unknown
Unknown
Laurasian
Laurasian
North American
Temperate (W)
Temperate, boreal
Temperate (W)
Unknown
Aquatic
Aquatic
Unknown
Diptera
Diptera
Unknown
Thorax
Thorax
Interstitial
Springs, riffles
Springs, riffles,
interstitial, pools
Crawling
Crawling
Crawling
Unknown
Diptera larvae
Unknown
Unknown
Unknown
Unknown
North American
Temperate
Unknown
Unknown
Unknown
Interstitial
Walking
Unknown
Unknown
North American
Temperate (E)
Unknown
Unknown
Unknown
Pools
Swimming
Unknown
Unknown
Gondwanan
Temperate (SW)
Unknown
Unknown
Unknown
Interstitial
Crawling
Unknown
Unknown
North American
Temperate, boreal
Aquatic
Diptera
Thorax
Springs, riffles, lakes
(profundal)
Crawling
Unknown
Unknown
Gondwanan
Pangean
Temperate
Temperate
Aquatic
Aquatic
Diptera
Diptera
Abdomen
Abdomen
Pools, lakes
Pools, lakes
Swimming
Swimming
Unknown
Unknown
Unknown
Unknown
Pangean
Throughout
Aquatic
Diptera,
odonata
Thorax, abdomen
Springs, riffles,
interstitial, pools,
lakes, temporary
pools
Walking,
swimming
Ostracoda,
Cladocera,
Copepoda,
Insect larvae
Copulation
649
650
Ian M. Smith, David R. Cook, Bruce P. Smith
III. OTHER ARACHNIDS IN
FRESHWATER HABITATS
A. Other Mites (ACARI)
1. Acariformes
a. Other Actinedida The mainly marine family Halacaridae is represented in both benthic and interstitial
freshwater habitats in North America by species of the
holarctic genera Porohalacarus, Lobohalacarus, Stygohalacarus, Soldanellonyx and Porolohmanella (Fig.
431). Halacarid mites have stilettoform chelicerae and
5-segmented pedipalps. Adults usually have four characteristic dorsal plates on the idiosoma and claws on
the tarsi of the legs. All instars have a sprawling gait
and crawl slowly on the substrate. Halacarid feeding
habits are generally poorly understood, but predaceous, parasitic and algivorous species are known.
Members of some species may be extremely abundant
in mats of vegetation and detritus in ponds and small
lakes. There are no comprehensive references for North
American freshwater halacarid mites. All known taxa
of freshwater Halacaridae were reviewed by K. H. Viets (1956) and K. O. Viets (1987) and species of the
German fauna, including all of the holarctic genera,
were treated by K.H. Viets (1936).
Homocaligidae is a small family of uncommon
species inhabiting wet vegetation and detritus at the
margin of ponds. Adults are well sclerotized and have
prominent hemispherical dorsal shields and long, protruding cheliceral stylets. These mites frequently venture beneath the surface, apparently aided by accessory
respiratory sacs and tubes. They have long legs and
crawl slowly on the substrate, but are thought to be
predaceous (Krantz 1978). Homocaligid taxa, including the North American species Homocaligus muscorum Habeeb, were reviewed by Wood (1969).
b. Oribatida The vast majority of oribatid mite
taxa are strictly terrestrial, but members of certain genera of the families Camisiidae, Trhypochthoniidae,
Malaconothridae, Hydrozetidae, Limnozetidae (Fig.
432), Ameronothridae, Tegeocranellidae and Zetomimidae are found only in freshwater habitats ranging from
springs and streams to large lakes. Oribatid mites have
chelate chelicerae and 5-segmented pedipalps. Adults
are variously sclerotized, and usually have distinctively
shaped prodorsal bothridia and claws on the tarsi of the
legs. All active instars of oribatid mites crawl slowly on
the substrate, feeding on both living and dead plant and
fungal material. The feeding habits of a species of Hydrozetes (Hydrozetidae) were investigated by Baker
(1985). Members of some species this genus may be
very numerous in mats of plant material and detritus.
Freshwater genera of oribatid mites can be identified
using the keys published by Balogh and Balogh (1992a,
b). Palearctic taxa, many of which are holarctic, were
treated by Balogh and Mahunka (1983). There is no
comprehensive reference for North American freshwater oribatid taxa, but systematic treatments have recently been published for the genera Mucronothrus
(Trhypochthoniidae) (Norton, Behan-Pelletier and
Wang, 1996), Limnozetes (Limnozetidae) (BehanPelletier, 1989) and Tegeocranellus (Tegeocranellidae)
(Behan-Pelletier, 1997).
c. Acaridida Most acaridid mites are terrestrial
saprophages, phytophages, fungivores or parasites, but
certain free-living species of the families Histiostomatidae and Hyadesiidae inhabit freshwater. Adult acaridids are generally soft-bodied, have 2-segmented pedipalps and lack both prodorsal trichobothria and claws
on the tarsi of the legs. Freshwater species are slowmoving crawlers typically associated with algal mats or
decaying detritus, although some have been reported
from water reservoirs in pitcher plants. Some species of
histiostomatid mites are reportedly predaceous on
microorganisms or oligochaete worms. Families of
freshwater Acaridida can be identified using the key
published by Krantz (1978).
2. Parasitiformes
Nearly all parasitiform taxa are strictly terrestrial,
but members of a few genera of the gamasid family Ascidae, subfamily Platyseiinae (Fig. 433), live exclusively
on wet plants and detritus and on the water surface of
marginal freshwater habitats. Platyseiinae have chelate
chelicerae and adults are long-legged mites with large
dorsal and ventral shields and conspicuous peritremes
laterally on the idiosoma. Active instars of these mites
walk about on the surface film, apparently feeding on
floating egg masses of nematocerous flies, especially
mosquitoes. Adult females of Platyseiinae regularly disperse between aquatic habitats as phoretic associates of
adult craneflies (Tipulidae).
B. Spiders
There are no truly aquatic spiders in North America, but many species of nurseryweb spiders (family
Pisauridae) are associated with aquatic habitats
(Carico, 1973). Species of Dolomedes (Fig. 434), the
genus with the greatest affinity for water, are large,
long-legged spiders with flattened, mottled bodies.
They are often called fishing spiders, dock spiders or
raft spiders because they venture onto water to hunt
and capture prey at or just below the surface film, and
can dive below the surface to escape potential preda-
16. Water Mites (Hydrachnida) and Other Arachnids
tors (Dondale and Redner, 1990). Members of some
species, such as Dolomedes tenebrosus, wander considerable distances from water (Carico, 1973; Dondale
and Redner, 1990), and even those of the most aquatic
North American species, Dolomedes triton, apparently
spend a significant part of their lives away from water
(Zimmermann and Spence, 1992, 1998).
The systematics and distribution of the nine described nearctic species of Dolomedes were thoroughly
reviewed by Carico (1973) and the five species occurring in Canada and Alaska were treated in detail by
Dondale and Redner (1990). Given the availability of
these relatively recent and comprehensive taxonomic
treatments, and in keeping with the level of coverage
for other groups in this chapter, we do not include a
key for the species of Dolomedes or the extensive morphological information that would be required to support it. The following brief summary of information is
intended as an introduction to the literature on the biology and ecology of these spiders.
Members of Dolomedes are typically found on
rocks or wood at the edge of water or on emergent or
floating vegetation. They move across the surface film
using a rowing gait and can dive beneath the surface to
remain submerged for as long as 30 min (Dondale and
Redner, 1990). When hunting, they typically remain
motionless with the anterior 2 – 3 pairs of legs resting
on the water surface. In this posture, they are able to
detect vibrations and chemical stimuli indicating the direction and proximity of potential prey (Roland and
Rovner, 1983). They apparently do not use visual stimuli to locate prey (Bleckmann and Lotz, 1987). Their
primary prey are adults of terrestrial and aquatic insects that have fallen on the water, including
Hemiptera, Diptera and Odonata (Zimmermann and
Spence, 1989), but they also capture and consume
small fish, tadpoles, and even small frogs, using their
venom to immobilize prey as much as four times their
own weight within a few minutes (Comstock, 1912;
Bleckmann and Lotz, 1987; Dondale and Redner,
1990). Cannibalism apparently accounts for up to 5%
of the prey of female D. triton (Zimmermann and
Spence, 1989, 1992, 1998).
Certain species of Dolomedes show habitat preferences that seem to be based on exposure and vegetation
type. For example, members of D. scriptus are most
common along larger, fast-flowing streams, with exposed rocky shorelines, those of D. vittatus prefer
edges of smaller and more shaded streams and individuals of D. triton and D. striatus frequent margins of
ponds, lakes, and calm backwaters of streams, including floating and emergent vegetation. The various
species have cryptic color patterns correlated with their
habitat preferences:those commonly found on rocks or
651
logs are predominately mottled with gray and brown
while those frequenting emergent vegetation have conspicuous longitudinal stripes (Carico, 1973; Dondale
and Redner, 1990).
Species of Dolomedes exhibit complex reproductive behavior. Females of Dolomedes triton mate only
once, and because males of this species cannot distinguish between mated and unmated females they are at
risk of sexual cannibalism. Males mature 5 – 10 days
before females, increasing their probability of finding
immature adult females and their likelihood of surviving multiple matings (Zimmermann and Spence, 1989,
1992). Males signal in the presence of females by waving the legs, drumming with the pedipalps, and twitching the body, to generate distinctive, regular, low-frequency waves in the water surface that differ from
those produced by prey (Roland and Rovner, 1983;
Bleckmann and Bender, 1987; Bleckmann and Lotz,
1987). They apparently also respond to pheromonal
cues, because their courtship displays can be induced
by water that has been in contact with conspecific females (Roland and Rovner, 1983). Sperm is transferred
to receptive females by inserting the embolus, a specialized intromittent organ at the tip of the male’s pedipalp, into a bursa copulatrix in the epigynum of the female (Carico, 1973; Roland and Rovner, 1983).
Gravid females of Dolomedes produce an egg sac
consisting of a silk-wrapped cluster of between 250
and 1500 eggs (Comstock, 1912; Carico, 1973; Zimmermann and Spence, 1992; Spence et al., 1996), and
carry it beneath their body using the chelicerae and a
silken dragline until the eggs are ready to hatch (Dondale and Redner, 1990). This parental care apparently
is necessary for successful hatching (Zimmermann and
Spence, 1992). Shortly before the eggs begin to hatch,
the female spins a nursery web, consisting of a tentlike
structure with the egg sac suspended within it (Dondale
and Redner, 1990). These webs are typically in low
plant growth or among rocks and are vigorously defended by females against potential predators. Newly
emerged juvenile spiders spend 3– 7 days in the web,
and disperse some time after their first molt either by
ballooning or walking (Zimmermann and Spence,
1992, 1998).
In central Alberta, Canada, members of
Dolomedes triton have approximately 12 juvenile instars but older individuals may experience supernumerary molts if exposed to warmer temperatures in autumn, possibly to delay maturation until spring.
Duration of instars 1 – 9 is the same for both sexes, but
duration of instars 10 – 12 is significantly longer for females, resulting in the protandry mentioned above
(Zimmermann and Spence, 1998). At this latitude, D.
triton is semivoltine, overwintering as instars 3 – 5 and
652
Ian M. Smith, David R. Cook, Bruce P. Smith
FIGURES 431 – 434 Scanning electron micrographs of adult mites. Fig. 431 Porolohmannella sp. (Halacaridae); Fig.
432 Hydrozetes sp. (Hydrozetidae); Fig. 433 Cheiroseius sp. (Ascidae). Fig. 434 Photograph of adult female
Dolomedes sp. (Pisauridae).
16. Water Mites (Hydrachnida) and Other Arachnids
9 – 11, with the first adults appearing in mid-May.
Males have an estimated average adult survival time of
9 – 13 days and disappear by the end of June, female
adults apparently live for 25 – 31 days and disappear by
late August or early September (Zimmermann and
Spence, 1992, 1998). Disappearance of males apparently reflects the influence of cannibalism by females
(Zimmermann and Spence, 1992). Overwintering is
facultative, with the later instars undergoing quiescence
based upon temperature control but the earlier instars
having dormancy determined by an interaction of photoperiod and temperature cues (Zimmermann and
Spence, 1998).
Cannibalism is an important cause of mortality in
Dolomedes populations. Interguild predation appears
to be a strong selective force on juveniles of D. triton,
and sexual cannibalism is correlated with the short seasonal occurrence of males of this species (Zimmermann
and Spence, 1989, 1992, 1998). Members of
Dolomedes are also eaten by birds (Carico, 1973) and
attacked by predatory pompilid and sphecoid wasps
(Carico, 1973; Dondale and Redner, 1990). Small- to
intermediate-sized juveniles of these spiders are eaten
by both water striders (Gerridae) and backswimmers
(Notonectidae) (Zimmermann and Spence, 1998). The
egg cocoons of Dolomedes are susceptible to infestation by hymenopterous parasitoids (Carico, 1973).
ACKNOWLEDGMENTS
We thank Michelle MacKenzie of the Biodiversity
Section of Agriculture and Agri-food Canada for her
expert help in assembling the plates of illustrations using computer-graphics software and for preparing
many of the water mite specimens examined during
preparation and testing of the keys. We also thank Drs.
Valerie Behan-Pelletier, Evert Lindquist and Charles
Dondale of the Biodiversity Section of Agriculture and
Agri-food Canada for providing illustrations and helpful comments for the sections dealing with oribatid
mites, ascid mites and spiders, respectively.
LITERATURE CITED
Åbro, A. 1979. Attachment and feeding devices of water-mite larvae
(Arrenurus spp.) parasitic on damselflies (Odonata, Zygoptera).
Zoologica Scripta 8:221 – 34.
Åbro, A. 1982. The effects of parasitic water mite larvae (Arrenurus
spp.) on zygopteran imagoes (Odonata). Journal of Invertebrate
Pathology 39:373 – 81.
Åbro, A. 1984. The initial stylostome formation by parasitic larvae
of the watermite genus Arrenurus on zygopteran imagines. Acarologia (Paris) 25:33 – 46.
653
Åbro, A. 1990. The impact of parasites in adult populations of Zygoptera. Odonatologica 19:223 – 234.
Alberti, G. 1977. Zur Feinstruktur und Funktion der Genitalnäpfe
von Hydrodroma despiciens (Hydrachellae, Acari). Zoomorphologie 87:155 – 164.
Alberti, G. 1979. Fine structure and probable function of genital
papillae and claparede organs of Actinotrichida. 501 – 507, in
Rodriguez, J. G., Ed.,. Recent Advances in Acarology, Vol. 2;
569 p.
Bader, C. 1938. Beitrag zur Kenntnis der Verdauungsvorgänge bei
Hydracarinen. Revue Suisse de Zoologie 45:721 – 806.
Bagge, P., Merilainen, J. J. 1985. The occurrence of water mites
(Acari: Hydrachnellae) in the estuary of the River Kyronjoki
(Bothnian Bay). Annales Zoologici Fennici 22:123 – 127.
Baker, G. T. 1985. Feeding, moulting and the internal anatomy
of Hydrozetes sp. (Oribatida: Hydrozetidae). Zoologische
Jahrbücher Anatomie 113:77 – 83.
Baker, G. T. 1996. Chemoreception in four species of water mites
(Acari: Hydrachnida): behavioural and morphological evidence.
Experimental and Applied Acarology 20:445 – 455.
Baker, R. L., Smith, B. P. 1997. Conflict between antipredator and
antiparasite behaviour in larval damselflies. Oecologia 109:
622 – 628.
Balogh, J., Balogh, P. 1992a. The oribatid mites genera of the World.
Vol. 1. Hungarian National Museum Press, Budapest, 263 p.
Balogh, J., Balogh, P. 1992b. The oribatid mites genera of the World.
Vol. 2. Hungarian National Museum Press, Budapest, 375 p.
Balogh, J., Mahunka, S. 1983. Primitive oribatids of the Palaearctic
region. Elsevier Scientific Publication, Amsterdam. 372 p.
Barr, D. W. 1972. The ejaculatory complex in water mites (Acari:Parasitengona): Morphology and potential value for systematics.
Life Sciences Contributions. Royal Ontario Museum 81; 87 p.
Barr, D. W. 1973. Methods for the collection, preservation, and study
of water mites (Acari: Parasitengona). Life Sciences Miscellaneous Publication. Royal Ontario Museum, 28 p.
Barr, D. W. 1979. Water mites (Acari, Parasitengona) sampled with
chemoluminescent bait in underwater traps. International Journal of Acarology, Vol. 5;187 – 194.
Barr, D. W. 1982. Comparative morphology of the genital acetabula
of aquatic mites (Acari, Prostigmata): Hydrachnoidea, Eylaoidea, Hydryphantoidea, Lebertioidea. Journal of Natural
History 16:147 – 160.
Behan-Pelletier, V. M. 1989. Limnozetes (Acari: Oribatida: Limnozetidae) of northeastern North America. Canadian Entomologist 121:453 – 506.
Behan-Pelletier, V. M. 1997. The semiaquatic genus Tegeocranellus
(Acari: Oribatida: Ameronothroidea) of North and Central
America. Canadian Entomologist 129:537 – 577.
Biesiadka, E. 1979. Wodopojki (Hydracarina) Pienin. Fragmenta
Faunistica 24:97 – 173.
Bleckmann, H., Bender, M. 1987. Water surface waves generated by
the male pisaurid spider Dolomedes triton Walckenaer during
courtship behavior. Journal of Arachnology 15:363 – 370.
Bleckmann, H., Lotz, T. 1987. The vertebrate-catching behaviour of
the fishing spider Dolomedes triton. Animal Behaviour
35:641 – 651.
Bohonak, A. J. 1998. Dispersal and gene flow in freshwater invertebrates. Ph.D. Thesis (unpublished), Cornell University, Ithaca,
NY.
Bonn, A., Gasse, M., Rolff, J., Martens, A. 1996. Increased fluctuating asymmetry in the damselfly Coenagrion puella is correlated
with ectoparasitic water mites: implications for fluctuating
asymmetry theory. Oecologia 108:596 – 598.
Böttger, K. 1962. Zur Biologie und Ethologie der einheimischen
Wassermilben Arrenurus (Megaluracarus) globator (Müll.),
654
Ian M. Smith, David R. Cook, Bruce P. Smith
1776, Piona nodata nodata (Müll.), 1776 und Eylais infundibulifera meriodionalis (Thon), 1989. (Hydrachnellae, Acari).
Zoologische Jahrbücher Systematik 89:501 – 584.
Böttger, K. 1965. Zur Ökologie und Fortpflanzungsbiologie von
Arrenurus valdiviensis Viets, K. O., 1964 (Hydrachnellae,
Acari). Zeitschrift für Morphologie und Ökologie der Tiere
55:115 – 141.
Böttger, K. 1970. Die Ernährungsweise der Wassermilben (Hydrachnellae, Acari). Internationale Revue der gesamten Hydrobiologie 55:895 – 912.
Böttger, K. 1972a. Vergleichend biologisch-ökologische Studien zum
Entwicklungszyklus der Süßwassermilben (Hydrachnellae,
Acari). I. Der Entwicklungszyklus von Hydrachna globosa und
Limnochares aquatica. Internationale Revue der gesamten Hydrobiologie 57:109 – 152.
Böttger, K. 1972b. Vergleichend biologisch-ökologische Studien zum
Entwicklungszyklus der Süßwassermilben (Hydrachnellae,
Acari). II. Der Entwicklungszyklus von Limnesia maculata und
Unionicola crassipes. Internationale Revue der gesamten Hydrobiologie 57:263 – 319.
Böttger, K., Schaller, F. 1961. Biologische und ethologische Beobachtungen an einheimischen Wassermilben. Zoologischer Anzeiger
167:46 – 50.
Carico, J. E. 1973. The nearctic species of the genus Dolomedes
(Araneae: Pisauridae). Bulletin of the Museum of Comparative
Zoology 144:435 – 488.
Chappuis, P. A. 1942. Eine neue Methode zur Untersuchung der
Grundwasserfauna. Acta Sciences, Mathematik und Natur,
Kolozsvar 6:3 – 7.
Cicolani, B., di Sabatino, A. 1991. Sensitivity of water mites to water
pollution. Modern Acarology 1:465 – 474.
Collins, N. C. 1975. Tactics of host exploitation by a thermophilic
water mite. Miscellaneous Publications of the Entomological
Society of America 9:359 – 370.
Collins, N. C., Mitchell, R. D., Wiegert, R. G. 1976. Functional
analysis of a thermal spring ecosystem, with an evaluation of
the role of consumers. Ecology 57:1221 – 1232.
Comstock, J. H. 1912. The spider book (revised and edited by
Gertsch, W. J., for the 1940 edition). Cornell University Press,
5th printing (1980), 729 p.
Cook, D. R. 1957. Order Acarina. Suborder Hydracarina. Genus
Protoarrenurus Cook, n. gen. in: Palmer, A. R., Ed. Miocene
arthropods from the Mojave Desert, California. Geological Survey Professional Paper (U.S.) No. 294-G, pp. 248 – 249.
Cook, D. R. 1974. Water mite genera and subgenera. Memoirs of the
American Entomological Institute 21;860 p.
Cook, D. R. 1980. Studies on Neotropical water mites. Memoirs of
the American Entomological Institute 31;645 p.
Cook, D. R. 1986. Water mites from Australia. Memoirs of the
American Entomological Institute 40;568 p.
Cook, D. R. 1988. Water mites from Chile. Memoirs of the American Entomological Institute 42;356 p.
Cook, D. R., Smith, I. M., Harvey, M. S. 2000. Assessment of lateral
compression of the idiosoma in adult water mites as a taxonomic character and reclassification of Frontipodopsis Walter,
Wettina Piersig and some other early derivative Hygrobatoidea
(Acari:Hydrachnida). Invertebrate Taxonomy 14:433 – 448.
Cook, W. J., Smith, B. P., Brooks, R. J. 1989. Allocation of reproductive effort in female Arrenurus spp. water mites (Acari:
Hydrachnidia; Arrenuridae). Oecologia 79:184 – 188.
Crowell, R. M. 1960. The taxonomy, distribution and developmental
stages of Ohio water mites. Bulletin of the Ohio Biological
Survey, (new series) 1;77 p.
Davids, C. 1973. The water mite Hydrachna conjecta Koenike, 1895
(Acari, Hydrachnellae), bionomics and relation to species of
Corixidae (Hemiptera). Netherlands Journal of Zoology
23:363 – 429.
Davids, C., Belier, R. 1979. Spermatophores and sperm transfer in
the water mite Hydrachna conjecta Koen. Reflections of the descent of water mites from terrestrial forms. Acarologia
21:84 – 90.
Davids, C., Heijnis, C. F., Weekenstroo, J. E. 1981. Habitat differentiation and feeding strategies in water mites in Lake Maarsseveen I. Hydrobiological Bulletin 15:87 – 91.
Dimock, R. V., Jr., LaRochelle, P. B. 1980. Chemically mediated host
recognition: a behavioral basis for the specificity of water mite
symbioses. American Zoologist 20:922.
Dimock, R. V., Jr., Davids, C. 1985. Spectral sensitivity and photobehavior of the water mite genus Unionicola. Journal of Experimental Biology 119:349 – 363.
Dondale, C. D., Redner, J. H. 1990. The wolf spiders, nurseryweb
spiders, and lynx spiders of Canada and Alaska (Araneae: Lycosidae, Pisauridae, and Oxyopidae). The insects and arachnids
of Canada; Part 17. Agriculture Canada. 383 pp.
Ellis-Adam, A. C., Davids, C. 1970. Oviposition and post-embryonic
development of the watermite Piona alpicola (Neuman, 1880).
Netherlands Journal of Zoology 20:122 – 137.
Elton, C. S. 1922. On the colours of water-mites. Proceedings of the
Zoological Society of London 82:1231 – 1239.
Eriksson, M. O. G., Henrikson, L., Oscarson, H. G. 1980.
Predator – prey relationships among water-mites (Hydracarina)
and other freshwater organisms. Archiv für Hydrobiologie
88:146 – 154.
Fairchild, W. L., O’Neill, M. C. A., Rosenberg, D. M. l987. Quantitative evaluation of the behavioural extraction of aquatic invertebrates from samples of sphagnum moss. Journal of the North
American Benthological Society 6:28l – 287.
Forbes, M. R. L. 1991. Ectoparasites and mating success in male
Enallagma ebrium damselflies (Odonata: Coenagrionidae).
Oikos 60:336 – 342.
Forbes, M. R. L., Baker, R. L. 1990. Susceptibility to parasitism: experiments with the damselfly Enallagma ebrium (Odonata: Coenagrionidae) and larval water mites, Arrenurus spp. (Acari: Arrenuridae). Oikos 58:61 – 66.
Forbes, M. R. L., Baker, R. L. 1991. Condition and fecundity of the
damselfly, Enallagma ebrium (Hagen): the importance of ectoparasites. Oecologia 86:335 – 341.
Garga, N. 1996. Aposematism in water mites (Acari: Hydracarina): a
predator defense mechanism, a phylogenetic hold-over and protection from damaging light. M.Sc. Thesis (unpublished),
Queens University, Kingston.
Gerecke, R., Schwoerbel, J. 1991. Water quality and water mites
(Acari: Actinedida) in the upper Danube region, 1959 – 1984.
Modern Acarology 1:483 – 491.
Gerecke, R., Smith, I. M., Cook, D. R. 1999. Three new species of
Apheviderulix gen. nov. and proposal of Apheviderulicidae
fam. nov. (Acari: Hydrachnida: Eylaoidea). Hydrobiologia
397:133 – 147.
Gledhill, T. 1982. Water-mites (Hydrachnellae, Limnohalacaridae,
Acari) from the interstitial habitat of riverine deposits in Scotland. Polish Archives of Hydrobiology 29:439 – 451.
Gledhill, T. 1985. Water mites — predators and parasites. Freshwater
Biological Association Annual Report 53:45 – 59.
Gliwicz, Z. M., Biesiadka, E. 1975. Pelagic water mites (Hydracarina) and their effect on the plankton community in a neotropical man-made lake. Archiv für Hydrobiologie 76:65 – 88.
Halik, L. 1955. O kopulach vodule Brachypoda versicolor (Müll.).
Biologia (Bratislava) 10:464 – 474.
Hevers, J. 1978. Zur Sexualbiologie der Gattung Unionicola (Hydrachnellae, Acari). Zoologische Jahrbücher Systematik 105:33 – 64.
16. Water Mites (Hydrachnida) and Other Arachnids
Hinton, H. E. 1964. Sperm transfer in insects and the evolution of
haemocoelic insemination. in Higham, K. C., (Ed.), Insect Reproduction, Symposium No. 2, Royal Entomological Society,
London, pp. 95 – 107.
Hungerford, H. B., Spangler, P. J., Walker, N. A. 1955. Subaquatic
light traps for insects and other animal organisms. Transactions
of the Kansas Academy of Sciences 58:387 – 407.
Imamura, T. 1952. Notes on the moulting of the adult of the water
mite, Arrenurus uchidai n. sp. Annotationes Zoologicae Japonenses 25:447 – 451.
Kerfoot, W. C. 1982. A question of taste: crypsis and warning
coloration in freshwater zooplankton communities. Ecology
63:538 – 554.
Knight, J. G. 1989. Investigation of reproductive biology, age, growth,
and feeding habits of the bayou darter, Ethiostoma rubrum
(Pisces, Percidae) from Bayou Pierre, Mississippi. M.Sc. Thesis
(unpublished), University of Southern Mississippi, Hattiesburg.
Koenike, F. 1891. Seltsame Begattung unter der Hydrachniden. Zoologischer Anzeiger 14:253 – 256.
Kouwets, F. A. C., Davids, C. 1984. The occurrence of chironomid
imagines in an area near Utrecht (the Netherlands), and their relations to water mite larvae. Archiv für Hydrobiologie
99:296 – 317.
Kowalik, W. 1981. Wodopojki (Hydracarina) rzek dorzecza Wieprza.
Annales Universitatis Mariae Curie-Sklodowska, Section C
36:327 – 352.
Kowalik, W., Biesiadka, E. 1981. Occurrence of water mites (Hydracarina) in the River Wieprz polluted with domestic-industry
sewage. Acta Hydrobiologica 23:331 – 347.
Krantz, G. W. 1978. A manual of Acarology. 2nd ed. Oregon State
University Press, Corvallis. 335 p.
Lanciani, C. A. 1971a. Host related size of parasitic water mites of
the genus Eylais. American Midland Naturalist 85:242 – 247.
Lanciani, C. A. 1971b. Host exploitation and synchronous development in a water mite parasite of the marsh treader Hydrometra
myrae (Hemiptera: Hydrometridae). Annals of the Entomological Society of America 64:1254 – 1259.
Lanciani, C. A. 1972. Mating behavior of water mites of the genus
Eylais. Acarologia 14:631 – 637.
Lanciani, C. A. 1976. Intraspecific competition in the parasitic water
mite, Hydryphantes tenuabilis. American Midland Naturalist
96:210 – 214.
Lanciani, C. A. 1978. Parasitism of Ceratopogonidae (Diptera) by the
water mite Tyrrellia circularis. Mosquito News 38:282 – 284.
Lanciani, C. A. 1979. The food of nymphal and adult water mites of
the species Hydryphantes tenuabilis. Acarologia 20:563 – 565.
Lanciani, C. A. 1980. Parasitism of the backswimmer Buenoa scimitra (Hemiptera: Notonectidae) by the water mite Hydrachna
virella (Acari:Hydrachnellae). Freshwater Biology 10:527 – 532.
Lanciani, C. A. 1982. Parasite-mediated reductions in the survival
and reproduction of the backswimmer Buenoa scimitra
(Hemiptera:Notonectidae). Parasitology 85:593 – 603.
Lanciani, C. A. 1983. Overview of the effects of water mite parasitism on aquatic insects. in Hoy, M. A., Cunningham, G. L.,
Knutson, L., Eds., Biological Control of Pests by Mites. Special
Publications of the University of California Agricultural Experimental Station 3304, pp. 86 – 90.
Lanciani, C. A. 1984. Crowding in the parasitic stage of the water
mite Hydrachna virella (Acari: Hydrachnidae). Journal of Parasitology 70:270 – 272.
Lanciani, C. A. 1986a. Competition among larval Tyrrellia circularis
(Acariformes: Limnesiidae) on the host Dasyhelea mutabilis
(Diptera: Ceratopogonidae). Florida Entomologist 69:438 – 439.
Lanciani, C. A. 1986b. Reduced survivorship in Dasyhelea mutabilis
(Diptera: Ceratopogonidae) parasitized by the water mite
655
Tyrrellia circularis (Acariformes: Limnesiidae). Journal of Parasitology 72:613 – 614.
Lanciani, C. A. 1987. Mortality in mite-infested, male Anopheles
crucians. Journal of the American Mosquito Control Association 3:107 – 108.
Lanciani, C. A. 1988. Sexual bias in host selection by parasitic mites
of the mosquito Anopheles crucians (Diptera: Culicidae). Journal of Parasitology 74:768 – 773.
Lanciani, C. A., Boyt, A. D. 1977. The effect of a parasitic water
mite, Arrenurus pseudotenuicollis (Acari: Hydrachnellae), on the
survival and reproduction of the mosquito Anopheles crucians
(Diptera: Culicidae). Journal of Medical Entomology 14:10 – 15.
Lanciani, C. A., May, P. G. 1982. Parasite-mediated reductions in the
growth of nymphal backswimmers. Parasitology 85:1 – 7.
Lanciani, C. A., McLaughlin, R. E. 1989. Parasitism of Coquillettidia
perturbans by two water mite species (Acari: Arrenuridae) in
Florida, U.S.A. Journal of the American Mosquito Control Association 5:428 – 431.
Lanciani, C. A., Smith, B. P. 1989. Constancy of stylostome form in
two water mite species. Canadian Entomologist 121:439 – 443.
Lang, P. 1905. Über den Bau der Hydrachnidenaugen. Zoologische
Jahrbücher Anatomie 21:453 – 494.
LaRochelle, P. B., Dimock, R. V. Jr. 1981. Behavioral aspects of host
recognition by the symbiotic water mite Unionicola formosa
(Acarina: Unionicolidae). Oecologia (Berlin) 48:257 – 259.
Leung, B., Forbes, M. R. L. 1997. Fluctuating asymmetry in relation
to indices of quality and fitness in the damselfly, Enallagma
ebrium (Hagen). Oecologia 110:472 – 477.
Lundblad, O. 1929. Über den Begattungsvorgang bei einigen
Arrhenurus-Arten. Zeitschrift für Morphologie und Ökologie
der Tiere 15:705 – 722.
Marshall, R. 1933. Water mites from Wyoming as fish food. Transactions of the American Microscopical Society. 52:34 – 41.
Marshall, R. 1940. On the occurrence of water mites in the food of
turtles. American Midland Naturalist 24:361 – 364.
Meyer, E. 1985. Der Entwicklungszyklus von Hydrodroma despiciens (Müller, O. F. 1776) (Acari: Hydrodromidae). Archiv
für Hydrobiologie, Supplement 66:321 – 453.
Meyer, E., Schwoerbel, J. 1981. Untersuchungen zur Phänologie der
Wassermilben (Hydracarina) des Mindelsees. Archiv für Hydrobiologie, Supplement 59:192 – 251.
Mitchell, R. D. 1955. Anatomy, life history, and evolution of the
mites parasitizing fresh-water mussels. Miscellaneous Publications of the Museum of Zoology, University of Michigan
89:1 – 41.
Mitchell, R. D. 1957a. Major evolutionary lines in water mites.
Systematic Zoology 6:137 – 148.
Mitchell, R. D. 1957b. Locomotor adaptations of the family
Hydryphantidae (Hydrachnellae, Acari). Abhandlungen herausgegeben vom naturwissenschaftlichen Verein zu Bremen
35:75 – 100.
Mitchell, R. D. 1957c. The mating behavior of pionid water mites.
American Midland Naturalist 58:360 – 366.
Mitchell, R. D. 1958. Sperm transfer in the water mite Hydryphantes
ruber Geer. American Midland Naturalist 60:156 – 158.
Mitchell, R. D. 1959. Life histories and larval behavior of arrenurid
water-mites parasitizing Odonata. Journal of the New York Entomological Society 67:1 – 12.
Mitchell, R. D. 1960. The evolution of thermophilous water mites.
Evolution 14:361 – 377.
Mitchell, R. D. 1961. Behavior of the larvae of Arrenurus fissicornis
Marshall, a water-mite parasitic on dragonflies. Animal Behaviour 9:220 – 224.
Mitchell, R. D. 1962. The structure and evolution of water mite
mouthparts. Journal of Morphology 110:41 – 59.
656
Ian M. Smith, David R. Cook, Bruce P. Smith
Mitchell, R. D. 1964a. A study of sympatry in the water mite genus
Arrenurus (family Arrenuridae). Ecology 45:546 – 558.
Mitchell, R. D. 1964b. An approach to the classification of water
mites. Acarologia 6:75 – 79.
Mitchell, R. D. 1966. Ecological aspects of sympatry in water mites.
Proceedings of the Japanese Society of Systematic Zoology
2:28 – 31.
Mitchell, R. D. 1969a. A model accounting for sympatry in water
mites. American Naturalist 103:331 – 346.
Mitchell, R. D. 1969b. Population regulation of larval water mites.
Proceedings of the 2nd International Congress of Acarology,
1967:99 – 102.
Mitchell, R. D. 1972. The tracheae of water mites. Journal of Morphology 136:327 – 335.
Modlin, R. F., Gannon, J. E. 1973. A contribution to the ecology and
distribution of aquatic Acari in the St. Lawrence Great Lakes.
Transactions of the American Microscopical Society 92:217 – 224.
Motas, C. 1928. Contribution à la connaissance des Hydracariens
français particilièrement du Sud-Est de la France. Travaux du
Laboratoire d’Hydrobiologie et Pisciculture. Université de
Grenoble 20:373 p.
Mullen, G. R. 1974. The taxonomy and bionomics of aquatic mites
(Acarina: Hydrachnellae) parasitic on mosquitoes in North
America. Ph.D. Thesis (unpublished), Cornell University, 263 p.
Mullen, G. R. 1975. Predation by water mites (Acarina: Hydrachnellae)
on immature stages of mosquitoes. Mosquito News 35:168 – 171.
Münchberg, P. 1935a. Über die bisher bei einigen Nematocerenfamilien (Culicidae, Chironomidae, Tipulidae) beobachteten ektoparasitaren Hydracarinenlarven. Zeitschrift für Morphologie
und Ökologie der Tiere 29:720 – 749.
Münchberg, P. 1935b. Zur Kenntnis der Odonatenparasiten, mit
ganz besonderer Berucksichtigung der Ökologie der in Europa
an Libellen schmarotzenden Wassermilbenlarven. Archiv für
Hydrobiologie 29:1 – 122.
Münchberg, P. 1937. Arrenurus planus Marsh. in USA. und A. papillator (O. F. Müll.) in der alten Welt, zwei ökologisch und morphologisch einander entsprechende Arten (Ordnung: Hydracarina). Archiv für Hydrobiologie 31:209 – 228.
Münchberg, P. 1938. Dritter Beitrag über die an Stechmücken
schmarotzenden Arrenurus-Larven (Ordn.: Hydracarina). Archiv
für Hydrobiologie 33:99 – 116.
Norton, R. A., Behan-Pelletier, V. M., Wang, H-f. 1996. The aquatic
oribatid mite genus Mucronothrus in Canada and western
U.S.A. (Acari: Trhypochthoniidae). Canadian Journal of Zoology 74:926 – 949.
Oliver, D. R., Smith, I. M. 1980. Host association of some pionid
water mite larvae (Acari: Prostigmata: Pionidae) parasitic on
chironomid imagos (Diptera: Chironomidae). Acta Universitatis
Carolinae, Biologica 1978:157 – 162.
Paterson, C. G. 1970. Water mites (Hydracarina) as predators of chironomid larvae (Insecta: Diptera). Canadian Journal of Zoology
48:610 – 614.
Pflügfelder, O. 1970. Schadwirkungen der Arrenurus-Larven (Acari,
Hydrachnellae) am Flügel der Libelle Sympetrum meridionale
Selys. Zeitschrift für Parasitenkunde 34:171 – 176.
Pieczynski, E. 1962. Notes on the use of light traps for water mites
(Hydracarina). Bulletin de l’Académie Polonaise des Sciences,
Classe II, Série des Sciences Biologiques 10:421 – 424.
Pieczynski, E. 1969. The trap method for ecological studies on water
mites (Hydracarina) in lakes. Proceedings of the 2nd International Congress of Acarology, 1967; pp. 103 – 106.
Pieczynski, E. 1976. Ecology of water mites (Hydracarina) in lakes.
Polish Ecological Studies 2:5 – 54.
Pieczynski, E., Kajak, Z. 1965. Investigations on the mobility of the
bottom fauna in the lakes Taltowisko, Mikolajskie and
Sniardwy. Bulletin de l’Académie Polonaise des Sciences, Classe
II, Série des Sciences Biologiques 13:345 – 353.
Poinar, G. O. 1985. Fossil evidence of insect parasitism by mites. International Journal of Acarology 11:37 – 38.
Prasad, V., Cook, D. R. 1972. The taxonomy of water mite larvae.
Memoirs of the American Entomological Institute 18:326 pp.
Proctor, H. C. 1989. Occurrence of protandry and a female-biased
sex-ratio in a sponge-associated water mite (Acari: Unionicolidae). Experimental and Applied Acarology 7:289 – 297.
Proctor, H. C. 1991a. The evolution of copulation in water mites: a
comparative test for nonreversing characters. Evolution 45:
558–567.
Proctor, H. C. 1991b. Courtship in the water mite Neumania papillator: males capitalize on female adaptations for predation. Animal Behaviour 42:589 – 598.
Proctor, H. C. 1992a. Sensory exploitation and the evolution of male
mating behaviour: a cladistic test using water mites (Acari: Parasitengona). Animal Behaviour 44:745 – 752.
Proctor, H. C. 1992b. Mating and spermatophore morphology of
water mites (Acari: Parsitengona). Zoological Journal of the
Linnaean Society 106:341 – 384.
Proctor, H. C. 1992c. Discord between field and laboratory sex
ratios of the water mite Neumania papillator Marshall (Acari:
Unionicolidae). Canadian Journal of Zoology 70:2483 – 2486.
Proctor, H. C. 1997. Sex ratios and chromosomes in water mites
(Acari: Parasitengona). Acarology IX 1:441 – 445.
Proctor, H. C., Pritchard, G. 1989. Neglected predators:water mites
(Acari: Parasitengona: Hydrachnellae) in freshwater communities.
Journal of the North American Benthological Society 8:100 – 111.
Proctor, H. C., Pritchard, G. 1990. Prey detection by the water mite
Unionicola crassipes (Acari: Unionicolidae). Freshwater Biology
23:271 – 279.
Proctor, H. C., Smith, B. P. 1994. Mating behaviour of the water
mite Arrenurus manubriator (Acari: Arrenuridae). Journal of
Zoology, London 232:473 – 483.
Punčochář, P., Hrbáček, J. 1991. Water mites in the plankton of
Hubenov reservoir and their relations to fish stock composition.
Modern Acarology 1:449 – 457.
Redmond, B. L., Hochberg, J. 1981. The stylostome of Arrenurus
spp. (Acari: Parasitengona) studied with the scanning electron
microscope. Journal of Parasitology 67:308 – 313.
Redmond, B. L., Lanciani, C. A. 1982. Attachment and engorgement
of a water mite, Hydrachna virella (Acari: Parasitengona), parasitic on Buenoa scimitra (Hemiptera: Notonectidae). Transactions of the American Microscopical Society 101:388 – 394.
Reilly, P., McCarthy, T. K. 1991. Watermite parasitism of Corixidae:
infection parameters, larval mite growth, competitive interaction and host response. Oikos 60:137 – 148.
Reinhardt, K. 1996. Negative effects of Arrenurus water mites on the
flight distances of the damselfly Nehallenia speciosa (Odonata:
Coenagrionidae). Aquatic Insects 18:233 – 240.
Riessen, H. P. 1982. Pelagic water mites: Their life history and seasonal distribution in the zooplankton community of a Canadian
lake. Archiv für Hydrobiologie, Supplement 62:410 – 439.
Robaux, P. 1974. Recherches sur le développement et la biologie des
acarines. “Thrombidiidee.” Mémories dee Museum National d’
Histoire Naturelle, Série A, Zoologie 85:1 – 186.
Roberts, E. A., Dimock, R. V., Jr., Forward, R. B., Jr. 1978. Positive
and host-induced negative phototaxis of the symbiotic water
mite Unionicola formosa. Biological Bulletin of the Marine
Laboratory (Woods Hole, Mass.) 155:599 – 607.
Robinson, J. V. 1983. Effects of water mite parasitism on the demographics of an adult population of Ischnura posita (Hagen)
(Odonata: Coenagrionidae). American Midland Naturalist
109:169 – 174.
16. Water Mites (Hydrachnida) and Other Arachnids
Roland, C., Rovner, J. S. 1983. Chemical and vibratory communication in the aquatic pisaurid spider Dolomedes triton (Araneae:
Pisauridae). Journal of Arachnology 11:77 – 85.
Rolff, J., Martens, A. 1997. Completing the life cycle: detachment of
an aquatic parasite (Arrenurus cuspidator, Hydrachnellae) from
an aerial host (Coenagrion puella, Odonata). Canadian Journal
of Zoology 75:655 – 659.
Rousch, J. M., Simmons, T. W., Kerans, B. L., Smith, B. P. 1997. Relative acute effects of low pH and high iron on the hatching and
survival of the water mite, Arrenurus manubriator and aquatic
insect, Chironomus riparius. Environmental Toxicology and
Chemistry 16:2144 – 2150.
Schmidt, U. 1935. Beiträge zur Anatomie und Histologie der Hydracarinen, besonders von Diplodontus despiciens O. F. Müller.
Zeitschrift für Morphogie und Ökologie der Tiere 30:99 – 175.
Schwoerbel, J. 1959. Ökologische und tiergeographische Untersuchungen über die Milben (Acari, Hydrachnellae) der Quellen
und Bäche des südlichen Schwarzwaldes und seiner Randgebiete. Archiv für Hydrobiologie, Supplement 24:385 – 546.
Simmons, T. W., Smith, I. M. 1984. Morphology of larvae, deutonymphs, and adults of the water mite Najadicola ingens
(Prostigmata: Parasitengona: Hygrobatoidea) with remarks on
phylogenetic relationships and revision of taxonomic placement
of Najadicolinae. Canadian Entomologist 116:691 – 701.
Smith, B. P. 1977. Water mite parasitism of water boatmen
(Hemiptera: Corixidae). M.Sc. Thesis (unpublished), University
of British Columbia, Vancouver, 117 pp.
Smith, B. P. 1983. The potential of mites as biological control agents
of mosquitoes. in Hoy, M. A., Cunningham, G. L., Knutson, L.
Eds. Biological Control of Pests by Mites. Special Publications
of the University of California Agricultural Experimental Station. No. 3304, pp. 79 – 85.
Smith, B. P. 1988. Host – parasite interaction and impact of larval water mites on insects. Annual Review of Entomology 33:487 – 507.
Smith, B. P. 1989. Impact of parasitism by larval Limnochares aquatica (Acari: Hydrachnidia; Limnocharidae) on juvenile Gerris
comatus, Gerris alacris, and Gerris buenoi (Insecta: Hemiptera;
Gerridae). Canadian Journal of Zoology 67:2238 – 2243.
Smith, B. P. 1998. Loss of larval parasitism in parasitengone mites.
Experimental and Applied Acarology 22:187 – 199.
Smith, B. P. 1999. Larval Hydrachnida and their hosts: biological inference and population structure. Pages 139–143 in: Needham,
G. R., Mitchell, R., Horn, D. J., Welbourn, W. C., Eds. Acarology IX: Vol. 2, Symposia, Ohio Biological Survey, Columbus,
OH, xvii 507 pp.
Smith, B. P., Cook, W. J. 1991. Negative covariance between larval
Arrenurus sp. and Limnochares americana (Acari: Hydrachnidia) on male Leucorrhinia frigida (Odonata: Libellulidae) and
its relationship to the host’s age. Canadian Journal of Zoology
69:226 – 231.
Smith, B. P., Laughland, L. A. 1990. Stimuli inducing detachment of
larval Arrenurus danbyensis (Hydrachnidia: Arrenuridae) from
adult Coquillettidia perturbans (Diptera: Culicidae). Experimental and Applied Acarology 9:51 – 62.
Smith, B. P., McIver, S. B. 1984a. The patterns of mosquito emergence
(Diptera: Culicidae; Aedes spp.): their influence on host selection
by parasitic mites (Acari: Arrenuridae; Arrenurus spp.). Canadian Journal of Zoology 62:1106 – 1113.
Smith, B. P., McIver, S. B. 1984b. Factors influencing host selection
and successful parasitism of Aedes spp. mosquitoes by Arrenurus spp. mites. Canadian Journal of Zoology 62:1114 – 1120.
Smith, B. P., McIver, S. B. 1984c. The impact of Arrenurus danbyensis Mullen (Acari: Prostigmata; Arrenuridae) on a population of
Coquillettidia perturbans (Walker) (Diptera: Cullcidae). Canadian Journal of Zoology 62:1121 – 1134.
657
Smith, I. M. 1976a. A study of the systematics of the water mite family Pionidae (Prostigmata: Parasitengona). Memoirs of the Entomological Society of Canada 98;249 pp.
Smith, I. M. 1976b. Paenecalyptonotus fontinalis n. gen., n. sp. with
remarks on the family Acalyptonotidae (Acari: Parasitengona:
Arrenuroidea). Canadian Entomologist 108:997 – 1000.
Smith, I. M. 1978. Descriptions and observations on host associations of some larval Arrenuroidea (Prostigmata: Parasitengona),
with comments on phylogeny in the superfamily. Canadian Entomologist 110:957 – 1001.
Smith, I. M. 1979. A review of water mites of the family Anisitsiellidae (Prostigmata: Lebertioidea) from North America. Canadian
Entomologist 111:529 – 550.
Smith, I. M. 1982. Larvae of water mites of the superfamily Lebertioidea (Prostigmata: Parasitengona) in North America with
comments on phylogeny and higher classification of the superfamily. Canadian Entomologist 114:901 – 990.
Smith, I. M. 1983a. Description of larvae of Neoacarus occidentalis
(Acari: Arrenuroidea: Neoacaridae). Canadian Entomologist
115:221 – 226.
Smith, I. M. 1983b. Description of adults of Neomideopsis siuslawensis n. gen., n. sp., with remarks on the family Mideopsidae (Acari: Parasitengona: Arrenuroidea). Canadian Entomologist 115:417 – 420.
Smith, I. M. 1983c. Description of Cowichania interstitialis n. gen.,
n. sp., and proposal of Cowichaniinae n. subfam., with remarks
on phylogeny and classification of Hydryphantidae (Acari:
Parasitengona: Hydryphantoidea). Canadian Entomologist 115:
523 – 527.
Smith, I. M. 1983d. Description of larvae and adults of Paramideopsis susanae n. gen., n. sp., with remarks on phylogeny and classification of Mideeopsidae (Acari: Parasitengona: Arrenuroidea).
Canadian Entomologist 115:529 – 538.
Smith, I. M. 1983e. Description of larvae of Nudomideopsis magnacetabula (Acari: Arrenuroidea: Mideopsidae) with new distributional records for the species and remarks on the classification
of Nudomideopsis. Canadian Entomologist 115:913 – 919.
Smith, I. M. 1983f. Descriptions of two new species of Acalyptonotus from western North America, with a new diagnosis of the
genus based on larvae and adults, and comments on phylogeny
and taxonomy of Acalyptonotidae (Acari: Parasitengona: Arrenuroidea). Canadian Entomologist 115:1395 – 1408.
Smith, I. M. 1983g. Descriptions of adults of a new species of Cyclothyas (Acari: Parasitengona: Hydryphantidae) from western
North America, with comments on phylogeny and distribution
of mites of the genus. Canadian Entomologist 115:1433 – 1436.
Smith, I. M. 1984. Larvae of water mites of some genera of Aturidae
(Prostigmata: Hygrobatoidea) in North America with comments
on phylogeny and classification of the family. Canadian Entomologist 116:307 – 374.
Smith, I. M. 1987. Water mites of peatlands and marshes in Canada.
Pages 3l – 46 in Rosenberg, D. M., Danks, H. V., Eds. Aquatic
insects of peatlands and marshes in Canada. Memoirs of the
Entomological Society of Canada l40; l74 pp.
Smith, I. M. 1989a. Description of deutonymphs and adults of Oregonacarus rivulicolus gen. nov., sp. nov. (Acari: Lebertioidea:
Anisitsiellidae). Canadian Entomologist 121:533 – 541.
Smith, I. M. 1989b. North American water mites of the family
Momoniidae Viets (Acari: Arrenuroidea). I. Description of adults
of Cyclomomonia andrewi gen. nov., sp. nov., and key to world
genera and subgenera. Canadian Entomologist 121:543 – 549.
Smith, I. M. 1989c. Description of two new species of Platyhydracarus gen. nov. from western North America, with remarks
on classification of Athienemanniidae (Acari: Parasitengona: Arrenuroidea). Canadian Entomologist 121:709 – 726.
658
Ian M. Smith, David R. Cook, Bruce P. Smith
Smith, I. M. 1989d. North American water mites of the family
Momoniidae Viets (Acari: Arrenuroidea). II. Revision of species of
Momonia Halbert, 1906. Canadian Entomologist 121:965 – 987.
Smith, I. M. 1989e. North American water mites of the family
Momoniidae Viets (Acari: Arrenuroidea). III. Revision of
species of Stygomomonia Szalay, 1943, subgenus Allomomonia
Cook, 1968. Canadian Entomologist 121:989 – 1025.
Smith, I. M. 1990a. Description of two new species of Stygameracarus gen. nov. from North America, and proposal of Stygameracarinae subfam. nov. (Acari: Arrenuroidea: Athienemanniidae). Canadian Entomologist 122:181 – 190.
Smith, I. M. 1990b. Proposal of Nudomideopsidae fam. nov. (Acari:
Arrenuroidea) with a review of North American taxa and description of a new subgenus and species of Nudomideopsis Szalay, 1945. Canadian Entomologist 122:229 – 252.
Smith, I. M. 1991a. Water mites (Acari: Parasitengona: Hydrachnida) of spring habitats in Canada. Pages 141 – 167 in:
Williams, D. D., Danks, H. V., Eds. Arthropods of springs, with
particular reference to Canada. Memoirs of the Entomological
Society of Canada 155; 217 pp.
Smith, I. M. 1991b. North American water mites of the genera
Phreatobrachypoda Cook and Bharatalbia Cook (Acari: Hygrobatoidea: Aturinae). Canadian Entomologist 123:465 – 499.
Smith, I. M. 1991c. North American water mites of the family
Momoniidae Viets (Acari: Arrenuroidea). IV. Revision of species
of Stygomomonia (sensu stricto) Szalay, 1943. Canadian Entomologist 123:501 – 558.
Smith, I. M. 1991d. Descriptions of new species representing new or
unreported genera of Lebertioidea (Acari: Hydrachnida) from
North America. Canadian Entomologist 123:811 – 825.
Smith, I. M. 1992a. North American species of the genus Chelomideopsis Romijn (Acari: Arrenuroidea: Athienemanniidae).
Canadian Entomologist 124:451 – 490.
Smith, I. M. 1992b. North American water mites of the family Chappuisididae Motas and Tanasachi (Acari: Arrenuroidea). Canadian Entomologist 124:637 – 723.
Smith, I. M. 1998. Chelohydracarus navarrensis gen. nov., sp. nov.
(Acari: Hydrachnida: Athienemanniidae). International Journal
of Acarology 24:327 – 330.
Smith, I. M., Cook, D. R. 1991. Water mites. Chapter 16, Pages
523 – 592 in Thorp, J., Covich, A., Eds. Ecology and classification of North American freshwater invertebrates. Academic
Press, San Diego, CA, 911 pp.
Smith, I. M., Cook, D. R. 1994. North American species of Neomamersinae Lundblad (Acari: Hydrachnida: Limnesiidae).
Canadian Entomologist 126:1131 – 1184.
Smith, I. M., Cook, D. R. 1996. New and unreported species of Neotyrrellia, Protolimnesia (Protolimnesella) and Centrolimnesia
(Acari: Hydrachnida: Limnesiidae) from the United States.
Anales del Instituto de Biología, Universidad Nacional
Autónoma de México, Serie Zoologia 67:265 – 277.
Smith, I. M., Cook, D. R. 1997. Description of Amoenacarus dixiensis gen. nov., sp. nov., and proposal of Amoenacaridae fam. nov.
(Acari: Hydrachnida: Arrenuroidea). International Journal of
Acarology 23:107 – 112.
Smith, I. M., Cook, D. R. 1998a. The Axonopsella-like water mites
of the United States (Acari: Hydrachnida: Aturidae). International Journal of Acarology. 24:311 – 325.
Smith, I. M., Cook, D. R. 1998b. Three new species of water mites
representing new genera of Thyadinae from North America
(Acari: Hydrachnida: Hydryphantidae). International Journal of
Acarology. 24:331 – 339.
Smith, I. M., Cook, D. R. 1999a. North American species of
Tartarothyas Viets (Acari: Hydrachnida: Hydryphantidae). International Journal of Acarology. 25:37 – 42.
Smith, I. M., Cook, D. R. 1999b. North American species of
Parathyasella stat. nov. and Amerothyasella gen. nov. (Acari:
Hydrachnida: Hydryphantidae). International Journal of Acarology. 25:43 – 50.
Smith, I. M., Cook, D. R. 1999c. An assessment of global distribution patterns in water mites (Acari: Hydrachnida). Pages
109 – 124 in: Needham, G. R., Mitchell, R., Horn, D. J.,
Welbourn, W. C., Eds. Acarology IX: Volume 2, Symposia.
Ohio Biological Survey, Columbus, OH, xvii 507 pp.
Smith, I. M., Oliver, D. R. 1976. The parasitic associations of larval
water mites with imaginal aquatic insects, especially Chironomidae. Canadian Entomologist 108:1427 – 1442.
Smith, I. M., Oliver, D. R. 1986. Review of parasitic associations of
larval water mites (Acari: Parasitengona: Hydrachnida) with insect hosts. Canadian Entomologist 118:407 – 472.
Sokolow, I. I. 1954. Khromosomnye kompleksy kleshchei i ikh
zhachenie dlia sistematiki i filogenii. Trudy Leningradskogo Obschchestva Estestvois-pytatelei 72:124 – 159.
Sokolow, I. I. 1977. The protective envelopes in the eggs of Hydrachnellae. Zoologischer Anzeiger 198:36 – 42.
Spence, J. R., Zimmermann, M., Wojcicki, J. P. 1996. Effects of food
limitation and sexual cannibalism on reproductive output of the
nursery web spider Dolomedes triton (Araneae: Pisauridae).
Oikos 75:273 – 382.
Stechmann, D.-H. 1980. Zum Wirtskreis syntopischer ArrenurusArten (Hydrachnellae, Acari) mit parasitischer Entwicklung an
Nematocera (Diptera). Zeitschrift für Parasitenkunde
62:267 – 283.
Steenbergen, H. A. 1993. Macrofauna-atlas of North-Holland: distribution maps and responses to environmental factors of aquatic
invertebrates. Cip-gegevens Koninkliijke Bibliotheek, Den
Haag, 651 pp.
Ten Winkel, E. H., Davids, C. 1985. Bioturbation by cyprinid fish
affecting the food availability for predatory water mites. Oecologia 67:218 – 219.
Ten Winkel, E. H., Davids, C. 1987. Chironomid larvae and their
food web relations in the littoral zone of Lake Maarsseveen.
Kaal Boek, Amsterdam. 145 pp.
Ten Winkel, E. H., Davids, C., de Nobel, J. G. 1989. Food and feeding strategies of water mites of the genus Hygrobates and the
impact of their predation on the larval population of the chironomid Cladotanytarsus mancus (Walker) in Lake Maarsseveen. Netherlands Journal of Zoology 39:246 – 263.
Uchida, T. 1940. Über die Begattungsakt bei Forelia variegator.
Annotationes Zoologicae Japonenses 19:280 – 282.
Vidrine, M. F. 1986. Revision of the Unionicolinae (Acari: Unionicolidae). International Journal of Acarology 12:233 – 243.
Viets, K. H. 1914. Über die Begattungsvorgange bei Acercus-Arten.
Internationale Revue gesamten Hydrobiologie und Hydrographie, Biological Supplement 6:1 – 10.
Viets, K. H. 1936. Wassermilben oder Hydracarina (Hydrachnellae und
Halacaridae). Die Tierwelt Deutschlands 31:1 – 288. 32:289 – 574.
Viets, K. H. 1956. Die Milben des Sußwassers und des Meeres
(Hydrachnellae und Halacaridae, Acari), Teil 2 / 3. Fischer
Verlag, Jena. 870 pp.
Viets, K. O. 1987. Die Milben des Sußwassers (Hydrachnellae und
Halacaridae [part.], Acari). 2. Katalog. Sonderbande Naturwissenschaftlichen Vereins Hamburg 8; 1012 pp.
Wainstein, B. A. 1980. Opredelitel lichinok vodjanych kleshchei.
(Key to larval water mites). Institut Biologii Vnutrennikh Vod,
Akademiia Nauk SSSR, 238 pp.
Weiberg, M, Edwards, D. D. 1997. Survival and reproductive output
of Chironomus tentans (Diptera: Chironomidae) in response to
parasitism by larval Unionicola foili (Acari: Unionicolidae).
Journal of Parasitology 83:173 – 175.
16. Water Mites (Hydrachnida) and Other Arachnids
Wesenburg-Lund, C. J. 1918. Contributions to the knowledge of the
postembryonal development of the Hydracarina. Videnskabelige
Meddelelser fra Dansk Naturhistorisk Forening 70:5 – 57.
Wiggins, G. B., Mackay, R. J., Smith, I. M. 1980. Evolutionary and
ecological strategies of animals in annual temporary pools.
Archiv für Hydrobiologie, Supplement 58:97 – 206.
Wiles, P. R. 1982. A note on the watermite Hydrodroma despiciens
feeding on chironomid egg masses. Freshwater Biology 12:83 – 87.
Wiles, P. R. 1984. Watermite respiratory systems. Acarologia
25:27 – 31.
Wood, T. G. 1969. The Homocaligidae, a new family of mites (Acari:
Raphignathoidea), including a description of a new species from
Malaya and the British Solomon Islands. Acarologia 11:711 – 729.
659
Zimmermann, M., Spence, J. R. 1989. Prey use of the fishing spider
Dolomedes triton (Pisuaridae: Araneae) an important predator
of the neuston community. Oecologia 80:187 – 194.
Zimmermann, M., Spence, J. R. 1992. Adult population dynamics
and reproductive effort of the fishing spider Dolomedes triton
(Araneae, Pisauridae) in central Alberta. Canadian Journal of
Zoology 70:2224 – 2233.
Zimmermann, M., Spence, J. R. 1998. Phenology and life-cycle regulation of the fishing spider Dolomedes triton Walckenaer
(Araneae, Pisauridae) in central Alberta. Canadian Journal of
Zoology 76:295 – 309.
This Page Intentionally Left Blank
17
DIVERSITY AND CLASSIFICATION
OF INSECTS AND COLLEMBOLA1
William L. Hilsenhoff
Department of Entomology
University of Wisconsin
Madison, Wisconsin 53706
I. Introduction
A. General Morphology
of Aquatic Insects
B. Common Techniques for
Studying Freshwater Insects
II. Aquatic Orders of Insects
A. Ephemeroptera — Mayflies
B. Odonata — Dragonflies
and Damselflies
C. Plecoptera — Stoneflies
D. Trichoptera — Caddisflies
E. Megaloptera — Fishflies and Alderflies
III. Partially Aquatic Orders of Insects
A. Aquatic Heteroptera — Aquatic
and Semiaquatic Bugs
B. Aquatic Neuroptera — Spongillaflies
C. Aquatic Lepidoptera — Aquatic
Caterpillars
D. Aquatic Coleoptera — Water Beetles
E. Aquatic Diptera — Flies and Midges
IV. Semiaquatic Collembola — Springtails
V. Identification of the Freshwater
Insects and Collembola
I. INTRODUCTION
This chapter includes orders and families in which
one or more life stages are truly aquatic and adapted
for survival under or on the water surface. Families
that live on or burrow into emergent aquatic vegetation, internal parasites of aquatic animals, and riparian
families that are closely associated with water but do
1
Much of the material on Collembola was contributed by Barbara L.
Peckarsky, Department of Entomology, Cornell University. The
author and editors greatly appreciate her generous contribution of
this material.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
A. Taxonomic Key to Orders
of Freshwater Insects
B. Taxonomic Key to Families of
Freshwater Ephemeroptera Larvae
C. Taxonomic Key to Families of
Freshwater Odonata Larvae
D. Taxonomic Key to Families of
Freshwater Plecoptera Larvae
E. Taxonomic Key to Families of
Freshwater Trichoptera Larvae
F. Taxonomic Key to Families of
Freshwater Megaloptera Larvae
G. Taxonomic Key to Adults of
Aquatic, Semiaquatic, and Riparian
Heteroptera
H. Taxonomic Key to Families of
Freshwater Coleoptera
I. Taxonomic Key to Families of
Freshwater Diptera Larvae
J. Taxonomic Key to Families of
Semiaquatic Collembola
Literature Cited
not inhabit it are not included. Terrestrial stages of
aquatic families are discussed briefly, but only as they
relate to the general life history of the family. In addition, brief mention is made of another arthropod
group, the semiaquatic springtails (class Entognatha,
order Collembola) (see Sections IV and V.J.)
There are 10 orders of insects that contain aquatic
species. Five of them (Ephemeroptera, Odonata, Plecoptera, Trichoptera, and Megaloptera) are “aquatic
orders” in which almost all species have aquatic larvae.
The remaining five orders (Heteroptera, Coleoptera,
Diptera, Lepidoptera, and Neuroptera) are “partially
661
662
W. L. Hilsenhoff
aquatic orders” in which most species are terrestrial,
but in which there are species or entire families that
have one or more life stages adapted for living in an
aquatic environment.
Three aquatic orders (Ephemeroptera, Odonata, and
Plecoptera) have a hemimetabolous life cycle, which includes three developmental stages: egg; larva; and adult.
The term “larva” is being used instead of “nymph” or
“naiad,” because in these orders the immature developmental stage is adapted for survival in aquatic environments and does not resemble the terrestrial adult, except
in Plecoptera. The other two aquatic orders (Trichoptera
and Megaloptera) have a holometabolous life cycle,
which includes four developmental stages: egg; larva;
pupa; and adult. In these orders the larva bears no resemblance to the adult.
Four of the five partially aquatic orders (Coleoptera, Diptera, Lepidoptera, and Neuroptera) also have
holometabolous life cycles and larvae that do not
resemble the adult. The fifth order, Heteroptera
(Hemiptera), has a paurometabolous life cycle, which
includes three developmental stages: egg, nymph,
and adult. In this order, nymphs inhabit the same habitat as adults and resemble them in many respects,
differing mostly by being smaller and less sclerotized,
having wing-pads instead of wings, and lacking genital
structures
FIGURE 1 Plecoptera larva (Perlidae); Acroneuria larva (dorsal)
showing various structures (thoracic gills and distal segments of legs
and cerci not shown).
A. General Morphology of Aquatic Insects
Morphological terms used for identification of
aquatic insects vary somewhat from order to order, but
several terms are commonly used for all orders. A Plecoptera larva (Figs. 1 and 2) is used to illustrate these
terms. Insects have three body regions: head, thorax,
and abdomen. The anterior region is the head, which
dorsally (Fig. 1) has a pair of antennae, a pair of compound eyes, and often three ocelli or simple eyes. On
the underside of the head are several mouthparts
(Fig. 2), including a labrum or upper lip, a pair of
mandibles, a pair of maxillae, and a labium or lower
lip. The mandibles usually are heavily sclerotized,
toothlike structures. The maxillae and labium are usually multisegmented and frequently have a pair of elongate, segmented structures called palps.
The thorax is immediately posterior to the head
and consists of three large segments (Fig. 1): a prothorax (anterior); a mesothorax; and a metathorax (posterior). The dorsal sclerite of each segment is referred to
as a notum, and the ventral sclerite as a sternum. Each
segment has a pair of legs that are divided into five
parts (Fig. 2): a coxa, trochanter, femur, tibia, and tarsus. The femur and tibia are usually elongate and the
tarsus has one-to-five segments, depending on the
FIGURE 2 Plecoptera larva (Perlidae); head and prothorax (ventral)
of Acroneuria larva showing mouthparts and structures of a leg (right
leg and distal segments of antennae not shown).
17. Diversity and Classification of Insects and Collembola
order. Each tarsus has one or two tarsal claws. Prefixes
pro-, meso- and meta-refer to the respective thoracic
segments. Thus, a pronotum would be on the prothorax and a mesofemur would be on the mesothorax. In
hemimetabolous and paurometabolous insect larvae or
nymphs, wing-pads (developing wings) are usually present dorsally on the meso- and metathorax.
The abdomen is the posterior body region and may
have as many as 10 visible segments, which are numbered posteriorly from the thorax. The dorsal sclerite
of each segment is called a tergum, the ventral sclerite a
sternum, and the lateral sclerite, when present, a pleuron. Terminal abdominal appendages vary widely or
may be absent; cerci are present in Plecoptera and several other orders. In some orders, gills of various types
may be present on the abdomen, thorax, or even on
the head.
A taxonomic key to orders of aquatic insects is
provided in Section V.A. Family level keys are also provided for aquatic larvae and aquatic adults in Sections
V.B – I.; keys to eggs, Heteroptera nymphs, pupae, and
terrestrial stages of aquatic species are not included.
The most recent keys to genera of aquatic insects in
North America appear in Merritt and Cummins
(1996). Regional keys to genera or species of aquatic
insects include those in Usinger (1956), Brigham et al.
(1982), Peckarsky et al. (1990), Clifford (1991), and
Hilsenhoff (1995c). The most recent keys to aquatic
stages of North American species are listed along with
regional species keys in tables that appear in the discussion of each order; reliable species keys do not exist for
many families and genera. Regional keys are important
because they are easier to use and may be more recent
than North American keys. Subsequent descriptions of
new species are not listed, but should be consulted
when species are being identified. References to recent
revisions and descriptions of new species may be found
in Merritt and Cummins (1996), bibliographies such as
those published annually by the North American Benthological Society, and through computer searches of
data bases such as Zoological Record and Dissertation
Abstracts Online.
B. Common Techniques for Studying
Freshwater Insects
1. Qualitative Sampling and Rearing
of Aquatic Insects
A D-frame or rectangular aquatic net with a mesh
size of 1 mm or less is recommended for sampling lotic
habitats. Shallow, rock or gravel riffles are sampled by
placing the net firmly against the bottom and disturbing
the substrate immediately upstream from the net with
663
one’s feet, allowing insects and debris washed into the
water to be carried into the net. This is commonly
called a “kick sample.” The contents of the net are
emptied into a shallow white pan containing 2 to 3 cm
of water, taking care not to collect so much debris that
it completely obscures the bottom of the pan. Insects
that swim or crawl from the debris are most easily collected with a curved forceps and placed into a collecting
jar containing 70% ethanol or Kahle’s fluid (15 parts
95% ethanol, 30 parts distilled water, 6 parts formalin,
and 1 part glacial acetic acid). The empty net should be
examined for insects that remain clinging to it.
Additional samples can be collected with a net by
pushing it under the stream bank or through silt and
debris in pools and rinsing excess silt from the net before placing the sample in the pan. It is also important
to remove pieces of decaying wood from the stream
and to collect insects that crawl from the wood as it
dries. Additional insects may be collected by examining
stones that are temporarily removed from the stream
bed. Deep areas of larger streams can be sampled by
using grabs or dredges. Ponar or Ekman grabs are most
commonly used, but the latter can be used only in soft
sediments.
Shallow lentic habitats, such as ponds, marshes,
swamps, and vegetated lake margins, are also best sampled with a flat-bottomed aquatic net. This is accomplished by pushing, pulling, or sweeping the net
through the vegetation or debris, rinsing excess silt
from the debris, and processing as described above.
Additional larger collections can be processed by placing them on a half-inch (13 mm) mesh screen placed
over a large white pan and allowing at least 15 min for
insects to crawl from the debris and drop into the pan.
This method is especially effective for collecting adult
Coleoptera and Heteroptera, as is the use of bottle
traps (Hilsenhoff, 1987). Wave-swept shorelines of
lakes can be sampled by disturbing the sand or gravel
bottom with one’s feet and immediately collecting suspended material with a net. Removal and examination
of decaying wood or stones from lake shores should
not be overlooked, because these substrates often harbor species that normally will not be collected with a
net. Deeper waters of lakes must be sampled with
grabs, dredges, or corers. Special habitats such as tree
holes or pitcher plants are best sampled with a large syringe or poultry baster. Appropriate sampling methods
are discussed under each order.
Since immature stages in many families of aquatic
insects cannot be identified to species, it is often necessary to rear them to the adult stage for identification.
This can be readily accomplished in most orders, but
the presence of a terrestrial pupal stage in Coleoptera,
Megaloptera, Neuroptera, and some Diptera makes
664
W. L. Hilsenhoff
rearing of larvae in these orders more difficult. Chapters on qualitative and quantitative sampling techniques and rearing procedures appear in Merritt and
Cummins (1996), Downing and Rigler (1984), McCafferty (1981), and Usinger (1956).
2. Preserving and Storing Aquatic Insects
Aquatic insects can most conveniently be killed
and preserved by placing them in a jar containing a liquid preservative. Many preservatives have been recommended and used, with 70 – 80% ethanol or isopropanol, or Kahle’s fluid being most popular. The
latter penetrates insect larvae more rapidly and preserves some colors better, but should be replaced with
70% ethanol after one or two days.
For permanent storage, 70% ethanol containing
3% glycerine is recommended, especially for larvae.
The glycerine serves to preserve larvae if the alcohol
evaporates due to a poorly fitting stopper. Most aquatic
entomologists store specimens in patent-lip vials with
neoprene stoppers. Screw-cap vials and cork-stoppered
shell vials should be avoided because of gradual evaporation of the alcohol, although screw-cap vials with
bevelled polyethyene inserts are satisfactory. One-dram
shell vials with polyethylene stoppers, — commonly
used for liquid scintillation studies, — are an inexpensive substitute for neoprene stoppered vials. Initially,
quality control of these polyethylene-stoppered vials
was poor and many leaked, but those purchased in recent years do not leak and retain fluid as well as neoprene-stoppered vials. They have a wider opening than
patent-lip or screw-cap vials, which facilitates removal
of insects. Because they are optically clear, insects that
are stored in these vials can be examined and often
identified under a microscope while still in the vial, if
the vial is only half-filled with alcohol so that a meniscus or bubble does not interfere with viewing. Adult
aquatic Coleoptera and Heteroptera are normally preserved on pins, but study collections can be preserved
in liquid, with several of the same species kept in a single vial to reduce storage space and curating time.
Adult Odonata are preserved on pins or in envelopes.
Adults of other orders also may be pinned, but are
most frequently preserved in alcohol. Storage in liquid
also facilitates removal of genitalia for study, which is
important for identification of many species.
For examination under a dissecting microscope,
specimens stored in liquid should be placed in a Syracuse watch glass or similar container and completely
covered with 95% alcohol. If 70% alcohol is used, insects may float and the alcohol will become cloudy as it
evaporates. If insects are not completely covered with
alcohol, a meniscus will cause distortions. A layer of
very small glass beads or silica sand in the bottom of
the watch glass permits insects to be imbedded for
viewing from any angle.
3. An Introduction to Biological Literature
A discussion of the general biology and habitat of
insects in each order and family is provided in Section
II, but it is often difficult to make generalizations at the
family level. Literature on biology of the numerous
genera and species is voluminous. References to biological literature pertaining to life histories and behavior
of species in each genus can be found in Merritt
and Cummins (1996) and in Current and Selected
Bibliographies on Benthic Biology that is published
each year by the North American Benthological
Society. Several review articles treat specific aspects of
biology such as drift of stream insects (Waters, 1972),
predator – prey relationships (Bay, 1974), detritus processing (Anderson and Sedell, 1979), and filter feeding
(Wallace and Merritt, 1980).
II. AQUATIC ORDERS OF INSECTS
A. Ephemeroptera — Mayflies
With only about 675 species in 20 families known
from North America, this is a relatively small
hemimetabolous order, but it is widespread and abundant in most regions. Larvae of all species are aquatic,
and while the great majority inhabit streams, several
inhabit a variety of permanent and temporary lentic
habitats. Ephemeroptera larvae can be found in
streams of all sizes and water temperatures, and even
occur in temporary streams. In lakes, they occur among
vegetation in littoral areas and sometimes also may be
found on the bottom in deep water or along waveswept shores. Frequently, they occur in vernal or permanent ponds and marshes. Larvae of the various families often differ so greatly from one another that they
can be easily identified to family in the field. They are
frequently an important source of food for fish in
streams. Because of the wide range of dissolved oxygen
requirements among the species, they are very important in biological monitoring of streams. Some species
that develop in lakes or large rivers may be sufficiently
abundant that emerging subimagos (pre-adults) and
adults create severe nuisance problems.
Except in warm waters of the southern United
States, most species are univoltine, with eggs either
hatching shortly after being laid and larvae developing
slowly over a 1-year period (slow seasonal life cycle),
or with eggs diapausing, followed by rapid development of the larvae after hatching several months later
(fast seasonal life cycle). Other species are bivoltine,
17. Diversity and Classification of Insects and Collembola
with rapid larval development and with eggs of the second generation diapausing over the winter. A few
species are multivoltine or semivoltine, with multivoltinism being especially prevalent in the South. The
number of larval instars varies widely among species,
and even within a species, with 12 or more having been
recorded for species that have been studied. After completion of larval development, the larva either crawls
from the water or swims or floats to the surface where
the subimago emerges. This winged subadult or “dun”
stage is unique among insects. Subimagos of most
species fly to nearby vegetation or supports where they
spend about a day before molting into the adult
(imago) stage. A few species molt almost immediately
to the adult, sometimes while in flight. Since most
subimagos are sexually mature, some females never
molt to the adult.
Adult mayflies do not feed, and generally live
only a few days. A few may live 2 weeks or more, and
some live only an hour or two. Mating normally takes
place when adult males form an aerial swarm to attract females and seize them when they enter the
swarm. In some species, adult males mate with
subimago females as they emerge from the water. Eggs
are laid mostly on the water surface by females that
repeatedly touch the water with their abdomen while
in flight or by females alighting briefly on the surface
to oviposit. Females of some species enter the water
on a substrate to oviposit on substrates beneath the
water. Most females die on the water surface upon
completion of oviposition.
Mayfly larvae can be readily distinguished from
other aquatic insects by their long, filamentous
caudal filaments and the presence of lateral or
ventrolateral gills on most of the first seven abdominal segments (Fig. 29). Usually three caudal filaments
are present (a pair of cerci and a median filament); in
a few species the median filament is so reduced that
only cerci are visible. Larvae most closely resemble
those of Plecoptera, which always have only cerci,
lack gills on the middle abdominal segments,
and have two tarsal claws (Figs. 44 and 45).
Ephemeroptera larvae have compound eyes, ocelli,
filamentous antennae, and 1-segmented tarsi with a
single claw (Figs. 9 and 29). Subimagos and adults
differ greatly from the larvae, having one or two
pairs of wings that they hold vertically above the
body when at rest, very large compound eyes, and no
gills or functional mouthparts. In the subimago, unmarked areas of the wings are opaque; in adults,
these areas are transparent.
While mayfly larvae mostly crawl on the substrate,
many are able and rapid swimmers. They swim by
moving their abdomen and caudal filaments up and
665
down rather than horizontally as stonefly do. Their
gills aid significantly in the uptake of oxygen, and
larvae of most species are capable of moving their gills
to increase water circulation. This enables larvae of
some species to inhabit lentic habitats or lotic habitats
that are less rich in oxygen. Almost all mayfly larvae
are herbivores or detritivores; larvae of a few species
are known to be predators on other invertebrates.
Subimagos and adults do not feed.
Larvae may be readily collected with a net by techniques already described. Those that inhabit deep water must be collected with grabs, and those in shallow
sand or silt areas can only be obtained by sieving these
substrates. Subimagos and adults may be collected by
sweeping stream-side vegetation, and large numbers are
often collected at lights.
Since the first edition (Thorp and Covich, 1991),
there have been several changes in the higher classification of Ephemeroptera, with the addition of five new
families (McCafferty, 1991). The order was divided
into three suborders by McCafferty (1991). Rectrachaeta, Setisura, and Pisciforma, which replace the former suborders Pannota and Schistonota. Although new
species and genera are still being discovered, recent revisions have often synonymized species, sometimes resulting in a reduction of the number of known species.
In general, larvae are poorly known and cannot be reliably identified to species in most families. Early books
by Needham et al. (1935), Burks (1953), and Berner
(1959) provided an essential background for more recent studies. The book by Edmunds et al. (1976) provides generic keys to adults and larvae as well as information on collecting, rearing, and mayfly biology;
generic keys to adults and larvae were updated by
G. F. Edmunds, Jr. and R. D. Waltz in Merritt and
Cummins (1996). The biogeography and evolution of
Ephemeroptera was reviewed by Edmunds (1972), and
Brittain (1982) reviewed the biology of this order. A
key to the families of freshwater Ephemeroptera larvae
is presented in Section V.B. and references to species
keys for identification of Ephemeroptera larvae are
provided in Table I. Information on the biology, habitat, and identification of each family is presented next.
Suborders and families are arranged alphabetically in
this and subsequent orders.
1. Families of Ephemeroptera — Suborder Pisciforma
a. Acanthametropodidae (2 Genera, 2 Species) This
family was formerly included in Siphlonuridae. The
large larvae inhabit sandy bottoms of medium-to-large
rivers where they feed primarily on larval Chironomidae. The rare and difficult-to-collect larvae are recognized by their bowed tibiae and tarsi and very long,
slender claws. One species occurs from Saskatchewan
666
W. L. Hilsenhoff
TABLE I Literature References to Species Keys
for Identification of Ephemeroptera Larvae
to Utah, the other from Wisconsin to South Carolina.
Life cycles are probably one year or longer.
Regional keys to larvae that include more than one family
California — Day (1956)
Florida — Berner and Pescador (1988)
Illinois — Burks (1953)
North and South Carolina — Unzicker and Carlson (1982)
United States — (Ephemeroidea) McCafferty (1975)
Keys to larvae listed by family
Ameletidae — (Alberta) Zloty and Pritchard (1997)
Ametropodidae — Allen and Edmunds (1976)
Baetidae
Acentrella — McCafferty et al. (1994)
Americabaetis — Waltz and McCafferty (1999)
Baetis, Acerpenna, Diphetor, Labiobaetis — Morihara
and McCafferty (1979); (Wisconsin) Bergman
and Hilsenhoff (1978)
Baetodes — McCafferty and Provonsha (1993)
Callibaetis — Check (1982)
Camelobaetidius — Lugo-Ortiz and McCafferty (1995);
Wieresma (1998)
Cloeodes — Waltz and McCafferty (1987a)
Fallceon — Lugo-Ortiz et al. (1994)
Labiobaetis — McCafferty and Waltz (1995)
Pseudocentroptiloides — Waltz and McCafferty (1989);
Wiersema and McCafferty (1998)
Baetiscidae — Pescador and Berner (1981); (Wisconsin)
Hilsenhoff (1984a)
Behningiidae — Hubbard (1994)
Caenidae
Brachycercus — (Northeast) Burian et al. (1997)
Brachycercus and Cercobrachys — Soldan (1984)
Caenis — Provonsha (1990)
Ephemerellidae
Attenella — Allen and Edmunds (1961b)
Caudatella — Allen and Edmunds (1961a)
Drunella — Allen and Edmunds (1962b)
Ephemerella — Allen and Edmunds (1965)
Eurylophella — Allen and Edmunds (1963b); Funk
and Sweeney (1994)
Serratella — Allen and Edmunds (1963a)
Timpanoga — Allen and Edmunds (1959); Allen and
Edmunds (1962a); McCafferty (1977)
Heptageniidae — (Wisconsin) Flowers and Hilsenhoff (1975)
Heptagenia — (Rocky Mountains) Bednarik and Edmunds (1980)
Macdunnoa — Flowers (1982)
Stenonema — Bednarik and McCafferty (1979)
Isonychiidae — Kondratieff and Voshell (1984)
Leptohyphidae
Leptohyphes — Allen (1978); (southwestern United States)
Allen and Murvosh (1987)
Leptophlebiidae
Neochoroterpes — Henry (1993)
Paraleptophlebia — (Oregon) Lehmkuhl and Anderson (1971)
Thraulodes — Allen and Brusca (1978)
Traverella — Allen (1973)
Neoephemeridae — Berner (1956); Bae and McCafferty (1998)
Oligoneuriidae
Homoeoneuria — Pescador and Peters (1980)
Potamanthidae — Bae and McCafferty (1991)
b. Ameletidae (1 Genus, 32 Species) This family
also was formerly included in Siphlonuridae. Larvae
are excellent swimmers that inhabit small, rapid
streams where they are mostly found clinging to vegetation or debris. They are recognized by their small,
oval, lamellate gills that have a sclerotized mesal or
lateral band (Fig. 30), short antennae, and lack of posterolateral spines on the apical abdominal segment.
Life cycles are probably mostly univoltine, with larvae
overwintering, and emergence from February to September, depending on latitude.
c. Ametropodidae (1 Genus, 3 Species) Larvae are
relatively large and flat, with very long meso- and
metathoracic claws and shorter prothoracic claws with
long spines (Figs. 21 and 22). They partially burrow
into sand in eddies of medium-to-large streams in western and north-central North America, and are generally
rare. Larvae have a unique method for feeding on suspended algae and detritus, which they gather with their
prothoracic legs from a vortex created in a pit that they
construct in the sand. Their life cycle is 1 year, with a
relatively long emergence period in late spring or early
summer.
d. Baetidae (18 Genera, 146 Species) Larvae are
relatively small, active swimmers, with lamellate abdominal gills (Fig. 29). They differ from larvae of other
species with lamellate gills on most of the first seven
abdominal segments in having relatively long antennae
(mostly twice the width of head), usually no pronounced posterolateral spines on abdominal segment 9,
and a median caudal filament that is often shorter
than the cerci, or vestigial (Fig. 29). Baetidae is a widespread and abundant family that occurs in a variety of
streams, and also in permanent and temporary ponds
or littoral zones of lakes. Lotic species are sometimes
univoltine, but more frequently bivoltine; most species
overwinter as diapausing eggs. Emergence of various
cohorts and species occurs from early spring to late autumn. Lentic species are often multivoltine. Recent
studies of larvae (Waltz and McCafferty, 1987a – c,
1989; McCafferty et al., 1994; McCafferty and Waltz,
1995; Lugo-Ortiz and McCafferty, 1998; Lugo-Ortiz
et al., 1999) have led to changes in the placement of
species within genera. Northern populations of some
species are parthenogenetic.
e. Metretopodidae (2 Genera, 9 Species) Larvae
reach a relatively large size and have lamellate gills,
very long meso- and metatarsal claws (Fig. 21), and
17. Diversity and Classification of Insects and Collembola
shorter protarsal claws that are distinctively bifid (Fig.
20). They are excellent swimmers and may be found
along banks and among vegetation in small-to-large
streams and occasionally along shores of lakes. They
occur in Canada, the upper Midwest, and in the East.
Their life cycle is 1 year, with emergence in the spring
or summer, depending on the species.
f. Pseudironidae (1 Genus, 1 Species) This family
was formerly included in Heptageniidae. Larvae are apparently rare in sand bottoms of larger streams where
they prey on larvae of Chironomidae. Larvae have a
somewhat flattened head with dorsal eyes and antennae, as in Heptageniidae. They can be recognized by
their long slender legs and claws and their unique gill
lamellae (Fig. 19), which have a fibrilliform basal tuft
and a ventral fingerlike projection. Life cycles are probably univoltine.
g. Siphlonuridae (4 Genera, 26 Species) Most larvae are relatively large with lamellate gills (Fig. 28).
They most closely resemble larvae of Baetidae, but larvae are usually larger, have short antennae, and pronounced spines on the posterolateral angles of the
ninth abdominal segment. Larvae are mostly herbivores – detritivores, but those of some species include
insects in their diet. Larvae occur in vernal forest pools,
stream-side pools, marshes, and swamps. Those of
some species inhabit streams, but move into small
pools just before emergence. Most, if not all, species
are probably univoltine.
2. Families of Ephemeroptera — Suborder
Rectrachaeta
a. Baetiscidae (1 Genus, 12 Species) Larvae are
distinctive, having a mesonotal carapace that covers
most of the abdomen and all abdominal gills (Fig. 9).
They occur throughout eastern and central North
America, and are rare or absent in the West. Larvae
partially burrow into silty margins or eddies of streams
and clean lakes, especially where the primary substrate
is sand. All species probably are univoltine and overwinter as larvae. Larvae crawl from the water onto a
substrate to emerge, with emergence occurring in
spring or early summer.
b. Behningiidae (1 Genus, 1 Species) This family
occurs mostly in the extreme southeastern United States,
with a single record from northwestern Wisconsin. The
unusual larvae burrow in the clean shifting sands of
larger streams with a fairly rapid current. The relatively
large larvae are unique, possessing crowns of bristles
on the head and pronotum (Fig. 10), having ventrolateral filamentous gills, and lacking claws on their legs,
667
which are highly modified for digging. The life cycle is
2 years, with a very long diapausing egg stage followed
by slow larval growth and emergence in late April and
early May. Unlike most other mayflies, larvae are
predators, feeding mostly on larvae of Chironomidae.
c. Caenidae (4 Genera, 24 Species) The small larvae are easily recognized by their nearly square operculate gills that are not mesally fused (Fig. 14). They are
widespread and common in a wide variety of lotic and
lentic habitats, including streams of all sizes, spring
seeps, marshes, swamps, ponds, and lakes. Here, they
frequent the sediments and often are partially covered
with silt. Larvae are generally more tolerant of low levels of dissolved oxygen than those in any other mayfly
family. Life cycles are poorly known. In the North,
most species are probably univoltine with overlapping
cohorts; in the South, two or more generations per year
are likely. Subimagos emerge at the surface, fly to a
support, and molt immediately to adult. Adults, which
live only a few hours, swarm and mate shortly after
ecdysis.
d. Ephemerellidae (8 Genera, 98 Species) Larvae
are recognized by the absence of gills on abdominal
segment 2, and lamellate or operculate gills on segments 3- or 4 – 7 (Fig. 15). Larvae of most species inhabit clean streams, where they are often abundant in
leaf litter, eddies, or near the banks. Those of a few
species may persist in organically enriched streams,
and others may be found on wave-swept lake shores.
While most larvae are herbivores – detritivores, larvae
of a few species are omnivorous, also feeding on small
invertebrates. Most species are univoltine, with a
spring emergence period and a summer egg diapause,
or with emergence in summer and no egg diapause.
Subimagos emerge from the surface or from protruding substrates.
e. Ephemeridae (4 Genera, 17 Species) Larvae are
recognized by their relatively large size, filamentous
gills that extend upward over their back (Fig. 4),
mandibular tusks that curve upward and outward, and
metatibiae with an acutely pointed apex (Fig. 8). Larvae burrow into sand or eddies in riffle areas of smallto-medium-sized streams or inhabit silt bottoms of
medium-to-large streams; they also inhabit sandy or silt
substrates in relatively clean lakes. In lakes and large
rivers, they may occur in such tremendous numbers
that synchronized emergences of subimagos create severe nuisance problems. Most species are apparently
univoltine with emergence mostly in the summer, but
2-year life cycles are known to occur in Canada. Larvae are primarily filter feeders, which circulate water
668
W. L. Hilsenhoff
through their burrows; they also eat algae and detritus
on the substrate surface. Species previously included in
Palingeniidae are again included in this family.
f. Leptohyphidae (2 Genera, 44 Species) This family was previously named Tricorythidae. Larvae are recognized by their triangular (Fig. 11) or oval (Fig. 12)
operculate gills on abdominal segment 2, which are
well separated distally, and by the absence of fringed
margins on other gills as found in Caenidae and
Neoephemeridae. Larvae are widespread, inhabiting
detritus, silt, and gravel in streams of all sizes; larvae of
some species are quite resistant to lowered levels of dissolved oxygen. Life cycles are univoltine to multivoltine, with eggs often diapausing over winter and emergence occurring in spring and summer after rapid
development of each generation.
g. Leptophlebiidae (10 Genera, 82 Species) The
medium-sized larvae characteristically have gills on segments 2 – 6 that are forked, bilamellate and pointed
apically, or terminating in filaments (Figs. 25 – 27). The
gills differ markedly from the lamellate, operculate, or
fringed gills found in other families. Larvae inhabit
slow or fast currents in a variety of streams where they
are found among debris, on rocks or wood, or in
gravel. While predominantly lotic, larvae of at least one
species migrate upstream in spring and leave the stream
to enter adjacent ponds or marshes prior to emergence.
Most species are univoltine, with eggs diapausing over
winter or summer and adults emerging in spring, summer, or autumn; bivoltine and multivoltine life cycles
occur in the South.
h. Neoephemeridae (1 Genus, 4 Species) This relatively uncommon family is confined to the East. Larvae
have square operculate gills and resemble Caenidae,
but the gills are fused mesally (Fig. 13), larvae grow to
a larger size than Caenidae larvae, and have metathoracic wing-pads, which Caenidae larvae lack. Larvae
cling to vegetation, debris, or undersides of rocks in
slow to fairly rapid streams. They apparently are univoltine with a slow seasonal development and emergence in spring or early summer, depending on latitude.
i. Polymitarcyidae (4 Genera, 8 Species) Larvae in
this small family of burrowing mayflies differ from
other burrowing mayflies by having mandibular tusks
that are curved inward and downward and are covered
with small spines. Unlike Ephemeridae larvae, the
metatibiae are rounded apically (Fig. 7). They inhabit
medium-to-large streams and lakes, burrowing into silt,
clay, or silt – gravel substrates; frequently they occur in
riffles. Larvae circulate water through their burrows
and filter out algae and detritus upon which they feed;
they also graze on algae and detritus from the substrate. All species apparently are univoltine, with synchronized emergences during the summer months. In
some species there is a prolonged winter egg diapause.
j. Potamanthidae (1 Genus, 5 Species) While having mandibular tusks and filamentous gills like other
burrowing mayfly larvae, gills of Potamanthidae larvae
are horizontal instead of arched over the back, and the
legs also are spread more horizontally and not modified
for digging (Fig. 6). They occur in medium-to-large
streams where they most often sprawl on gravel and
sand in shallow runs. They have a 1-year life cycle with
emergence during the summer months.
3. Families of Ephemeroptera — Suborder Setisura
a. Arthropleidae (1 Genus, 1 Species) This family
was formerly included in Heptageniidae. Larvae have
dorsal antennae and eyes and were previously placed in
the family Heptageniidae, but they differ in having extremely long maxillary palps that curve around both
sides of the head and have long setae (Fig. 18). Larvae
inhabit shallow ponds, bogs, or channels with almost
no flow. Eggs hatch in spring and larvae develop
rapidly with emergence in late spring. The life cycle is
univoltine with a long egg diapause. Larvae use their
long maxillary palps to strain plankton from the water
for food. They are generally uncommon, occurring in
northeastern North America.
b. Heptageniidae (14 Genera, 135 Species) The distinctive larvae are readily identified by their flattened
appearance and the dorsal location of their eyes and
antennae (Fig. 16). Larvae in some genera lack the median caudal filament. They are widespread and abundant in streams; some also occur on wave-swept shorelines of lakes. Larvae inhabit rocks, wood, debris, and
other substrates to which they cling. Most species are
probably univoltine, but a few may be bivoltine, especially in the South. Emergence occurs from spring to
autumn, depending on the species, with most emerging
in the summer.
c. Isonychiidae (1 Genus, 17 Species) This family
was formerly included in Oligoneuriidae. The large larvae are strong swimmers and fairly common in rapidly
flowing streams where they cling to rocks or debris.
They are recognized by the double row of long setae on
the inner margins of the profemurs and protibiae
(Fig. 23), which they use to gather algae and small
insects from the current. They also have gill tufts at
the base of the procoxae (Fig. 23) and bilamellate gills
with a fibrilliform tuft, which separate them from
17. Diversity and Classification of Insects and Collembola
Oligoneuriidae. Life cycles are mostly bivoltine or
univoltine, depending on the latitude.
d. Oligoneuriidae 2 Genera, 8 Species) Larvae have
two rows of long setae on the inner margin of the prothoracic legs, which distinguish them from larvae of
other mayflies, except those of Isonychiidae. Gills are
small and either rounded or elongate (Fig. 24), quite
unlike in larvae of Isonychiidae. They use the setae on
their prothoracic legs to filter diatoms and other algae
from the current. Larvae are found either in sandy bottoms of streams or on sticks or rocks. Depending on
latitude and species, life cycles are univoltine to multivoltine with emergence during the summer months.
B. Odonata — Dragonflies and Damselfies
This relatively small hemimetabolous order has
two distinctive suborders in North America:
Anisoptera (dragonflies), and Zygoptera (damselflies).
Only about 434 species are known from North America. Larvae of all species are aquatic, with about twothirds of them living in lentic habits and one-third in
lotic environments. Larvae of lotic species occur in all
types of permanent stream habitats, including gravel
and rock riffles, debris along banks, bank vegetation,
soft sediments, and sand. Those of lentic species inhabit permanent and temporary ponds, marshes,
swamps, and littoral and shoreline areas of lakes. Because all larvae attain a relatively large size and are
predators, mostly on other insects, they are often important apex predators in the invertebrate community.
Adults are also predators, feeding on mosquitoes and
other flying insects, and thus Odonata are regarded as
highly beneficial insects.
Life cycles are relatively long, mostly 1 year in Zygoptera, and 1 – 6 years in Anisoptera. Slow seasonal
developmental cycles predominate, but fast seasonal
cycles occur in both suborders. There are normally 11
or 12 larval instars, but the number may vary somewhat between species or even within a species. When
larval development is complete, the larva crawls from
the water onto emergent vegetation or onto the bank,
where ecdysis occurs. Larvae of some species may
crawl several meters from the shore to emerge. Adults
may live for several weeks. Newly emerged “teneral”
adults are feeble fliers for a day or two and vulnerable
to predation. After the teneral stage has passed, adults
often disperse widely into fields and other habitats to
feed for a week or more on flying insects. They then return to breeding sites where males patrol selected areas,
chasing away rival males and other species. Before mating, the male transfers sperm from primary genitalia at
the tip of the abdomen to a secondary genital organ on
669
the venter of abdominal segment 2. Using claspers on
the terminal segment, a male will seize a female of the
same species that flies into his territory, grasping her either by the pronotum (Zygoptera) or by the head
(Anisoptera). The female then loops the tip of her abdomen forward and upward to receive sperm from the
penis on the second abdominal sternum of the male.
Eggs are laid singly in plant tissues (endophytic
oviposition) above, at, or below the water surface, or
they may be laid singly or in masses on plants, debris,
or other substrates, or randomly on the water surface
(exophytic oviposition). Oviposition is endophytic in all
Zygoptera and in Aeshnidae and Petaluridae; it is exophytic in other families. In many species, especially
species of Zygoptera and Libellulidae, the male continues to grasp the female while she is ovipositing, protecting her from other males. Eggs hatch after an incubation period of one to several weeks, or after a long
diapause. The first larval instar is a prolarval stage that
lasts only a few minutes to several hours. Prolarvae lack
the well-developed appendages of other larval instars.
Odonata larvae can be readily distinguished from
all other aquatic insects by their elongate, hinged
labium that has been modified for seizing and grasping
prey (Figs. 32 – 34). They have conspicuous compound
eyes and relatively short filamentous antennae (Fig.
43). In Zygoptera, the abdomen is elongate and thin,
terminating in three long, vertically oriented, caudal
lamellae (Fig. 31). In Anisoptera, the abdomen is relatively broad and terminates in three triangular-shaped,
pointed appendages (Fig. 44), the upper one being
called the “epiproct” and the lower ones “paraprocts.”
In between them are triangular-shaped cerci, which
usually are distinctly shorter than the epiproct or paraprocts. Adults do not resemble larvae, being much
more elongate, having conspicuous genitalia, two pairs
of elongate net-veined wings, and much larger compound eyes. Like Ephemeroptera, the primitive wings
of Odonata cannot be folded. Wings of Anisoptera are
held horizontally to the side when at rest; those of Zygoptera are held vertically just above or slightly to the
side of the abdomen.
Odonata larvae move about on the substrates by
crawling; some actively stalk prey. Zygoptera larvae
may swim by moving their abdomen and caudal lamellae from side to side. Anisoptera larvae can move
rapidly when disturbed by expelling water from a rectal chamber. There are tracheal gills within the rectal
chamber, and movement of water in, and out of, the
chamber aids in respiration of Anisoptera larvae. In
Zygoptera, respiration is greatly enhanced by the caudal lamellae, enabling larvae of most species to live in
water with relatively low levels of dissolved oxygen.
Most larvae can be easily collected with an aquatic net
670
W. L. Hilsenhoff
using techniques described earlier. Adults are collected
with large aerial nets after being sighted. Zygoptera
and the Anisoptera adults that alight on vegetation are
relatively easy to capture, but many large Anisoptera
adults are extremely difficult to capture with a net.
Most larvae of North American Odonata can be
identified to species by using keys and descriptions
provided by Walker (1953, 1958) and Walker and
Corbet (1975), Needham and Westfall (1955), and
Westfall and May (1996). Most keys to Anisoptera larvae, however, are old, and problems often arise when
using them. A revised edition of Dragonflies of North
America (Needham et al., 2000) will contain new
larval and adult keys. Because Odonata are relatively
homogeneous, differences between families, genera,
and species are not as obvious as in most other orders.
Larvae of Corduliidae cannot be readily separated
from those of Libellulidae without identifying them to
genus. There is also considerable disagreement about
the status of various genera among odonatologists as
well as some disagreement about family status. But, at
the species level, North American Odonata are better
known as larvae and adults than aquatic insects in any
other order. This is perhaps because they have attracted the interest of many amateur odonatologists
who have made significant contributions to our knowledge. Books by Corbet (1962) and Miller (1987), and
a review article by Corbet (1980), summarize knowledge about the biology of dragonflies. References to
species keys for the identification of Odonata larvae
are provided in Table II, and a key to families of freshwater Odonata appears in Section V.C. Information on
the biology, habitat, and identification of dragonfly
and damselfly larvae in each family is presented in the
following.
1. Families of Odonata — Suborder
Anisoptera (Dragonflies)
a. Aeshnidae (11 Genera, 39 Species) Larvae are
large, with elongate tapered abdomens and can be distinguished from other Anisoptera by their elongate
shape, flat prementum, and thin six- or seven-segmented antennae (Fig. 37). Most inhabit lentic habitats, especially weedy permanent ponds, marshes, and
littoral areas of lakes. Here, they move about on the
vegetation and stalk prey composed of invertebrates
and small vertebrates. Larvae of a few species are lotic
and inhabit either slower areas of streams or riffles.
Life cycles of 2 – 4 years are likely for most species, especially in the North. One species, Anax junius, which
does not overwinter in the North, migrates south in autumn and returns in early spring to oviposit. Larvae of
this species develop more rapidly than other Aeshnidae
and are bivoltine in the South, but rarely complete two
TABLE II Literature References to Species Keys
for Identification of Odonata Larvae
Regional keys to larvae that include more than one family
Canada and Alaska — (Zygoptera) Walker (1953); (Anisoptera)
Walker (1958); (Anisoptera) Walker and Corbet (1975)
California — Smith and Pritchard (1956)
Florida — (Zygoptera) Daigle (1991b); (Anisoptera) Daigle (1992)
North America — (Anisoptera) Needham and Westfall (1955);
Needham et al. (2000); (Zygoptera) Westfall and May (1996)
North and South Carolina — Huggins and Brigham (1982)
Southeastern United States — (lotic Anisoptera) Louton (1982)
Texas — Young and Bayer (1979)
Utah — (Anisoptera) Musser (1962)
Keys to larvae listed by family
Aeshnidae
Gomphaeschna — Dunkle (1977)
Coenagrionidae
Enallagma — (West Coast) Garrison (1984)
Ischnura — (California) Garrison (1981)
Corduliidae
Somatochlora — Daigle (1991a)
Gomphidae
Dromogomphus — Westfall and Tennessen (1979)
Lanthus — Carle (1980)
Ophiogomphus — Carle (1992)
Progomphus — Tennesson (1993)
Libellulidae
Ladona — Bennefield (1965)
Sympetrum — Tai (1967)
generations in the northern United States. Flight periods of most species are during the summer months.
b. Cordulegastridae (1 Genus, 8 Species) The large,
hairy, somewhat elongate larvae with small anterolateral eyes can be easily distinguished from other
Anisoptera by the jagged teeth on the palpal lobes of
their spoon-shaped prementum (Fig. 39). They inhabit
small streams, where they lie partially concealed in silt
to ambush prey, mostly at the upstream edge of pools.
A 3 – 5-year life cycle is likely for most species, with
adults flying from spring into early summer. Eggs are
laid in the substrate of shallow riffles.
c. Corduliidae (7 Genera, 49 Species) Larvae are
short and broad, with a spoon-shaped prementum, and
cannot be readily separated from those of Libellulidae.
In most, the palpal lobes are more deeply scalloped
(Figs. 40 and 41) than in Libellulidae (Fig. 42), but
larvae in one genus of Libellulidae also have deeply
scalloped palpal lobes. Larvae are mostly lentic, with
2 – 4-year life cycles probably predominating in the
North and shorter cycles in the South. They commonly
occur in marshes, swamps, cool ponds, and littoral areas of lakes. However, several lotic species have larvae
that inhabit debris in streams of all sizes. Synchronized
17. Diversity and Classification of Insects and Collembola
emergences may lead to formation of exceptionally
large swarms. Adult flight periods are mostly from
May into early July.
d. Gomphidae (15 Genera, 97 Species) As in larvae
of Aeshnidae and Petaluridae, the prementum is flat,
but gomphid larvae have relatively shorter and broader
abdomens. Their broad, four-segmented antennae are
diagnostic (Fig. 36). Larvae of most species inhabit
streams, where they may be found in riffles or partially
buried in silt or sand in slower reaches. Those of other
species inhabit lentic habitats, especially small permanent ponds and littoral areas of lakes, where they lie
concealed in the substrate to ambush prey. Life cycles
are long, usually 2 – 4 years, depending on water temperature and food. Most species have flight periods in
late spring and summer; in some species eggs may diapause over the winter.
e. Libellulidae (26 Genera, 105 Species) The short,
broad larvae closely resemble those of Corduliidae, but
in almost all genera the crenulations on the palpal lobe
of the mentum are extremely shallow (Fig. 42). Larvae
occur in a variety of permanent and temporary lentic
habitats where they crawl about in the vegetation and
debris; occasionally they are found in vegetation along
margins of streams. Littoral areas of lakes, permanent
ponds, vernal ponds and marshes, cattail marshes,
sphagnum swamps, and bogs all are habitats for larvae
of a great variety of species. Many species are univoltine, with some having eggs that diapause. Other
species may require two or more years to complete larval development. Flight periods range from spring
through autumn, depending on the species.
f. Macromiidae (2 Genera, 9 Species) Larvae are
readily recognized by their relatively broad abdomens,
extremcly long legs, and the presence of a frontal horn
that projects forward between the eyes (Fig. 43). They
are found among debris in the current of medium-tolarge streams or in margins of lakes exposed to wave
action. Life cycles of 2 – 4 years are likely. Flight periods extend from March through August, depending on
the species.
g. Petaluridae (2 Genera, 2 Species) Both North
American species are rare; neither has been found in
the central part of the continent. Larvae of one semiterrestrial species inhabit bog seepages at high elevations
in the western mountains; those of the other species
have been found in bogs and seeps in eastern forests.
The relatively broad larvae have a flat prementum, relatively broad six-or seven-segmented antennae that are
hairy (Fig. 38), and patches of black bristles on the
671
posterior abdominal terga. Because of their cold habitat, up to 6 years may be required to complete larval
development. The flight period ranges from early
spring in Florida to summer farther north.
2. Families of Odonata — Suborder
Zygoptera (Damselflies)
a. Calopterygidae (2 Genera, 8 Species) The large
larvae are readily distinguished from other Zygoptera
by their very elongate first antennal segment (Fig. 31).
All species are lotic, the larvae inhabiting permanent
streams of all sizes, where they are found mostly
among bank vegetation and accumulations of debris.
Species are univoltine or semivoltine, with flight periods from late May into October.
b. Coenagrionidae (13 Genera, 96 Species) The relatively small larvae differ from those of Calopterygidae
in having a short basal antennal segment, and differ
from those of Lestidae in not having the base of the
prementum greatly narrowed (Fig. 34). They are predominantly lentic, although species in one genus
(Argia) inhabit riffles of streams and a few species in
other genera occur along banks of streams. Larvae of
lentic species are found mostly in permanent ponds,
marshes, swamps, and littoral areas of lakes; occasionally, they occur among vegetation in parts of streams
with little or no current. They crawl about on the vegetation and ambush or stalk prey, which consists mostly
of small invertebrates. Most species are univoltine,
some having eggs that diapause over the winter; a few
species are bivoltine. Adults fly from spring to autumn,
depending on the species.
c. Lestidae (2 Genera, 18 Species) The long, thin
larvae with their long, parallel-sided caudal lamellae
are distinctive. They differ from all other Zygoptera in
having a prementum that is greatly narrowed and elongate basally (Fig. 33). Larvae of one genus (Archilestes)
with 2 species are lotic. Larvae of other species (Lestes)
are lentic, only occasionally being found among emergent vegetation along slow streams. They commonly
inhabit vegetation in both permanent and vernal ponds
and marshes and have a 1-year life cycle. Eggs are laid
in vegetation above the water and often diapause until
autumn or the following spring. Eggs hatch when the
now dead vegetation in which they were laid falls into
the water, or in vernal ponds, when water floods the
dead vegetation. Adults fly from late spring through
summer.
d. Protoneuridae (2 Genera, 2 Species) This tropical family is found only in Texas where larvae inhabit
streams. Larvae are similar to those of Coenagrionidae,
672
W. L. Hilsenhoff
but their caudal lamellae are thick in the basal half and
thin and leaflike in the distal half (Fig. 35).
C. Plecoptera — Stoneflies
In this relatively small hemimetabolous order at
least 614 species are known from North America, and
it is likely that many additional species will be described as the taxonomy of various families and genera
is studied in greater detail and mountain streams are
collected more thoroughly. The order is poorly represented in middle North America; species richness is
greatest in mountain areas where fast, cold streams
abound. Larvae of all species are aquatic. Almost all of
them inhabit streams, but those of a few species have
adapted to living in cold oligotrophic lakes. In the
West, larvae of species in at least two genera inhabit
the hyporheic zone of certain streams to depths of 10
m or more.
Stonefly larvae are an important part of the stream
ecosystem. They provide food for fish and invertebrate
predators, and often are top predators in the invertebrate food chain. They are not sufficiently abundant to
create nuisance problems, and because larvae of most
species are intolerant of lowered levels of dissolved
oxygen they play an important role in biological monitoring of streams.
Most stoneflies are univoltine, but several larger
species require 2 or 3 years to complete development.
Species with 2 or 3 year life cycles in the North may be
univoltine in the South. Univoltine species often have
fast seasonal life cycles, with eggs or small larvae diapausing during the summer and rapidly developing in
autumn, winter or spring. Many species, however, have
a slow seasonal life cycle with eggs hatching within a
few weeks and larval development progressing slowly
throughout the year. Species that have a slow seasonal
life cycle in the North may have a bivoltine, fast seasonal life cycle in the South.
The number of larval instars varies with the species
and even among individuals within a species, with
12 – 23 or more instars having been recorded for
species that have been studied. After completing development, larvae crawl from the water onto a convenient
substrate where ecdysis occurs. Adults then crawl or fly
to nearby vegetation or other structures to find a mate.
In most species this is accomplished by drumming with
their abdomen; males initiate the drumming signal and
females respond. After repeated drumming and crawling, males find a female and mate. When eggs have matured, the female extrudes them into a mass on the tip
of her abdomen. Females of some species fly over the
water and land briefly on its surface to deposit their
eggs; others jettison their egg mass while flying above
the water. In still other species, females crawl into the
water on a protruding substrate to oviposit beneath the
surface. In most species the female will lay multiple egg
masses.
Stonefly larvae can be distinguished from most
other aquatic insects by their two long filamentous
cerci and generally elongate or flattened appearance
(Figs. 45, 48, 49, 54, 56, and 58). They have relatively
long antennae, distinct compound eyes, two or three
ocelli, chewing mouthparts, two pairs of thoracic wingpads, and three-segmented tarsi with two claws on
each tarsus (Figs. 1 and 2). They are likely to be confused only with Ephemeroptera larvae, which also have
long cerci, but Ephemeroptera larvae usually have three
caudal filaments instead of two, have only one tarsal
claw, and have gills on the middle abdominal segments.
Gills on Plecoptera larvae, if present, are on the head,
thorax, and / or basal or apical segments of the abdomen. Adult stoneflies resemble their larvae, the most
noticeable difference being the presence of two pairs of
net-veined wings that fold flat over the abdomen; a few
species are brachypterous or apterous. Genitalia are
well developed in adults and gills are reduced to remnants in species that have them as larvae.
Stonefly larvae generally crawl on the substrate;
when forced to swim, most do so weakly by moving
their abdomen from side to side. Oxygen for respiration is obtained from the water through the cuticle,
with thoracic and, sometimes, abdominal gills aiding
oxygen uptake in most larger species. Because they generally lack extensive gills, most Plecoptera larvae occur
only in oxygen-rich water. Larval feeding habits vary
among families and also among species within a family
or genus. As in many other aquatic insects, larval feeding habits within a species will vary with developmental stage and availability of food.
Adult stoneflies are relatively short-lived; most live
a few days to 2 weeks, some a little longer. Several
species feed as adults, mostly on algae, lichens, and riparian leaves or flowers. Larvae can be readily collected with an aquatic net by using standard sampling
techniques. Adults of most species can be collected by
using a net to sweep bank vegetation or by examining
rocks, tree trunks, and bridges in the vicinity of
streams. Adults of some species, however, move high
into trees and can be collected only by using nets with
very long handles or by using sticky traps. Several
species of adults are attracted to lights, while others are
rarely found at lights.
Through the years there have been many changes
in the higher classification of Plecoptera, but in the last
two decades it has remained quite stable. The order is
divided into two suborders, Euholognatha and Systellognatha. The former incudes four families (Capniidae,
17. Diversity and Classification of Insects and Collembola
Leuctridae, Nemouridae, and Taeniopterygidae) that
previously were in the family Nemouridae; larvae in all
of these families are primarily herbivores – detritivores.
Larvae of two families of Systellognatha (Perlidae and
Chloroperlidae) are predators, in two other families
(Peltoperlidae and Pteronarcyidae) they are herbivores – detritivores, and in the remaining family
(Perlodidae) feeding habits vary with the species and
larval age.
The biology of Plecoptera was reviewed by Hynes
(1976) and a recent book by Stewart and Stark (1988)
discusses classification, phylogeny, biogeography, ecology, and behavior of Plecoptera. This book includes
keys and illustrations for larvae in all North American
genera, notes on their biology, and the known distribution of North American species. Species in some genera
can be identified as mature larvae as well as adults; in
other genera identification of larvae is not possible because larvae of several species remain unknown. About
55% of North American Plecoptera species are known
in the larval stage. Information on the biology, habitat,
and identification of larvae in each family is presented
next. References to species keys for the identification of
stonefly larvae are provided in Table III and a key to
families of freshwater Plecoptera larvae appears in Section V.D.
1. Families of Plecoptera – Suborder Euholognatha
a. Capniidae (10 Genera, 151 Species) Capniidae
are “winter stoneflies.” Larvae are generally small,
dark colored, and elongate (Fig. 56). They closely resemble larvae of Leuctridae, but can be recognized by
their generally shorter and broader metathoracic wingpads, which are nearly as widely separated as the
mesothoracic wind-pads (Fig. 56), and by posterior abdominal terga that are usually widened and fringed
posteriorly. Larvae inhabit streams of all sizes. They
are especially abundant in smaller streams and spring
seeps; some even inhabit streams that become dry in
the summer.
Larvae in two genera occur in cold oligotrophic
lakes. Life cycles are univoltine, with emergence in late
winter or early spring. Eggs hatch a few weeks after being laid, and after some development, larvae of most
species diapause. Larval development resumes in autumn, with some species developing most rapidly in
late autumn and winter. A few species have a slow seasonal life cycle with steady development from late
spring through autumn. Larvae are mostly detritivores
that feed on allochthonous debris, especially decaying
leaves. In the North, several species may emerge on
warm days while streams are still ice covered; adults
often can be found crawling on the snow. Adults of
some species are apterous or brachypterous.
673
TABLE III Literature References to Species Keys
for Identification of Plecoptera Larvae
Regional keys to larvae that include more than one family
British Columbia (southwest) — Ricker (1943)
Louisiana — Stewart et al. (1976)
Minnesota — Harden and Mickel (1952)
North and South Carolina — Unzicker and McCaskill (1982)
Northeastern North America — Hitchcock (1974)
Ozark and Ouachita mountains — Poulton and Stewart (1991)
Pennsylvania — Surdick and Kim (1976)
Rocky Mountains — Baumann et al. (1977)
Saskatchewan — Dosdall and Lehmkuhl (1979)
Texas — Szczytko and Stewart (1977)
Keys to larvae listed by family
Capniidae — (eastern Canada) Harper and Hynes (1971b)
Leuctridae — (eastern Canada) Harper and Hynes (1971a)
Nemouridae — (eastern Canada) Harper and Hynes (1971d)
Peltoperlidae
Soliperla — Stark (1983)
Perlidae
Agnetina — Stark (1986)
Hesperoperla — Baumann and Stark (1980)
Paragnetina — Stark and Szczytko (1981)
Perlinella — Kondratieff et al. (1988)
Perlodidae — (Wisconsin) Hilsenhoff and Billmyer (1973)
Diploperla — Kondratieff et al. (1981)
Helopicus — Stark and Ray (1983)
Hydroperla — Ray and Stark (1981); Ricker (1952)
Isogenoides — Ricker (1952)
Isoperla — (western North America) Szczytko and
Stewart (1979)
Malirekus — Stark and Szczytko (1988)
Setvena — Stewart and Stanger (1985)
Pteronarcyidae — (West Virginia) Tarter et al. (1975)
Taeniopterygidae — Harper and Hynes (1971c)
Taeniopteryx — Fullington and Stewart (1980)
b. Leuctridae (7 Genera, 55 Species) The relatively
small, elongate larvae are similar to those of Capniidae,
but have metathoracic wing-pads that are similar in
shape, but distinctly closer together than those on the
mesothorax (Fig. 58). Larvae also resemble those of
Chloroperlidae, which have shorter cerci and meso-and
metathoracic wing-pads that are equally far apart (Fig.
49). Larvae are found mostly among gravel in small,
cool, permanent streams where they feed on decaying
allochthonous material; one genus occurs in temporary
streams. Most species are univoltine with emergence
from spring to autumn, depending on the species. A
semivoltine life cycle occurs in at least one northern
species.
c. Nemouridae (12 Genera, 72 Species) The small,
relatively dark-colored larvae are short and broad, with
divergent metathoracic wing-pads in more mature
larvae and metathoracic legs that reach or exceed the
apex of the abdomen (Fig. 54). Most nemourids are
674
W. L. Hilsenhoff
univoltine, with emergence from spring to autumn, depending on the species. Fast seasonal life cycles predominate, with an extended egg diapause in some species.
At least one species is semivoltine. Larvae are detrital
feeders that inhabit debris and soft sediments in smaller
streams and spring seeps; they are often the only stonefly larvae in spring seeps and runs. Larvae in at least
one genus inhabit temporary streams, and those of
another species have been collected from Arctic lakes.
d. Taeniopterygidae (6 Genera, 35 Species) Like
Capniidae, these are univoltine “winter stoneflies” that
emerge in late winter or very early spring and probably
spend the summer as diapausing larvae. In size and
shape, larvae rsemble those of Perlodidae (Fig. 48),
having divergent metathoracic wingpads and elongate
abdomens, but they are not as heavily patterned as
most Perlodidae, they move much more slowly, and
they swim very inefficiently. Larvae are herbivores – detritivores that occur in permanent streams of all sizes.
They are frequently found among bank vegetation.
2. Families of Plecoptera — Suborder Systellognatha
a. Chloroperlidae (13 Genera, 77 Species) Larvae
are generally small, elongate, rounded in cross section,
and have short cerci (Fig. 49). They are similar to larvae
of Capniidae and Leuctridae and are difficult to distinguish from the larvae of Leuctridae in the field. They
are not likely to be confused with Capniidae, however,
because late-instar larvae of Chloroperlidae are present
in late spring and summer, while those of Capniidae occur in late autumn, winter, and early spring. Larval
mouthparts (Fig. 46) differ greatly from those of Capniidae and Leuctridae (Fig. 47), reflecting the carnivorous
feeding habits of Chloroperlidae in later larval instars.
Larvae of Chloroperlidae inhabit mostly gravel bottoms
of cold, small- to medium-sized streams. Life histories
are poorly known. Univoltine and semivoltine fast seasonal life cycles have been documented, with emergence
in late spring or early summer.
b. Peltoperlidae (6 Genera, 20 Species) The broad,
flat, roachlike appearance of the medium-sized larvae is
distinctive (Fig. 45). They are often abundant in mountain streams of the eastern, southern, and western United
States, but are absent from central North America.
Larvae inhabit smaller streams, where they feed on decaying leaves, detritus, and associated microorganisms.
Many species are semivoltine; some may be univoltine.
Adults emerge in late spring or early summer.
c. Perlidae (15 Genera, 72 Species) Larvae are relatively large, broad, stoneflies (Fig. 1) that are readily
recognized by the presence of branched filamentous
gills at the base of each leg (Fig. 2) and the absence of
gills on abdominal sterna. They are widespread and
common, inhabiting permanent streams of all sizes
where they prey on other macroinvertebrates. They are
often the dominant insect predators, although earlyinstar larvae may feed on detritus and algae. Most larvae are found in fast water, especially among debris in
riffles. Species in most genera have a 2 or 3 year slow
seasonal life cycle in northern regions; univoltine life
cycles may occur in the South. One common genus is
univoltine with a fast seasonal life cycle. Emergence in
most species is during the summer months.
d. Perlodidae (30 Genera, 122 Species) Larvae are
mostly small- to medium-sized stoneflies, which usually
have distinctive patterns dorsally. They lack the thoracic
gills of the similarly shaped and patterned larvae of Perlidae. The metathoracic wing-pads are divergent and the
abdomen is rather elongate (Fig. 48). While larvae of
some species of Perlodidae are herbivores – detritivores
throughout their larval development, most species are
predominantly predators. They develop in all sizes and
types of streams; some even develop in cold oligotrophic
lakes. All species are apparently univoltine, with emergence in spring. Eggs of many species diapause during
the summer and hatch in autumn, but in other species
eggs hatch within a few weeks after being laid.
e. Pteronarcyidae (2 Genera, 10 Species) These
large stoneflies require 1 – 4 years to complete development. They have a slow seasonal life cycle, with univoltine life cycles occurring only in the south. Larvae
are elongate, and can be readily recognized by the
branched filamentous gills at the base of thoracic legs
and on the first two or three abdominal sterna. They
feed mostly on coarse particulate organic matter and
inhabit wood or accumulations of debris in small- to
medium-sized permanent streams with moderate-torapid currents. Emergence occurs in spring or early
summer, being earliest in the south.
D. Trichoptera – Caddisflies
Trichoptera is a rather large holometabolous order.
Larvae and pupae of all species are aquatic, with one
or two exceptions, and represent an important part of
the insect fauna of most streams. About 1400 species
are known from North America. Most larvae occur in
lotic habitats, but those of some species in half of the
families inhabit lentic habitats. Adults are terrestrial,
mothlike insects, that sometimes are attracted to lights
in large numbers. Most have very long filiform antennae, and their wings, which they fold rooflike over
their body, are covered with hairlike setae. Eggs of
17. Diversity and Classification of Insects and Collembola
most species are laid in masses in the water; adults of a
few species lay their eggs on vegetation above the water
or in moist soil of temporary ponds that will eventually
be flooded. Most species are univoltine, often with
rather synchronized emergences; however, in some
species different cohorts emerge throughout the warm
weather period. A few species are bivoltine, with the
tendency for bivoltinism increasing farther south, and
several species, especially those that develop in coldwater streams, are known to be semivoltine, especially
in the northern part of their range.
Adult caddisflies can be identified to species by
using the many keys and descriptions that have been
published, but larval taxonomy lags, with about half of
the described species still unknown in the larval stage.
This makes accurate identification of larvae to species
impossible in most genera, and larvae of four genera
remain unknown. Wiggins (1996) published excellent
keys, descriptions, and illustrations for larvae of
known genera. Ross (1944) developed species keys for
larvae in many genera that occur in the central United
States as well as keys to adults, but in most of these
genera there are species for which larvae are unknown,
so his larval keys must be used with caution.
Trichoptera is divided into three suborders, Annulipalpia, Integripalpia, and Spicipalpia (Wiggins and
Wichard, 1989; Frania and Wiggins, 1995). This parallels the division of the order into three superfamilies as
used by Wiggins (1977) and others, and to some extent
the classification proposed by Weaver and Morse
(1986). In the Annulipalpia, larvae use silk from their
labial glands to construct retreats and nets, which they
use to filter or gather food such as algae, detritus, or
macroinvertebrates. Larvae pupate within their retreat
in a silk cocoon that has small openings at each end
through which water circulates. In the suborder Spicipalpia, larvae in the four families are either free-living
or construct portable cases that are barrel-shaped,
purselike, or saddle-shaped. All larvae pupate in a completely closed cocoon of parchmentlike silk, differing
from other caddisflies that have openings in their pupal
case to allow circulation of water. Free-living larvae
(Rhyacophilidae and Hydrobiosidae) are mostly predators on other arthropods. Larvae that build cases
(Glossosomatiae and Hydroptilidae) are mostly herbivores that feed on periphyton. In the suborder Integripalpia, all larvae use silk from their labial glands to
construct portable tubular cases of plant or mineral
materials. Cases vary widely in size and shape, and are
often sufficiently distinctive to be employed in generic
and species identification. Integripalpia larvae are
mostly herbivores or detritivores, but in some families (Brachycentridae, Leptoceridae, Limnephilidae,
Molannidae, Odontoceridae, and Phryganeidae) omni-
675
vores or predators can be found. Larvae in eight families of Integripalpia have a prosternal horn, that larvae
in other families lack. All larvae pupate within their
last larval case after it has been attached to a substrate
and closed at both ends. Small openings at each end
permit water circulation.
Trichoptera larvae have a pair of simple eyes,
chewing mouthparts, antennae that are so short that
they often are difficult to see, three pairs of thoracic
legs with a single tarsal segment and tarsal claw, and a
pair of fleshy prolegs on the last abdominal segment
(Figs. 66 and 67). Many larvae have single or branched
gills on the abdominal segments. Most species have five
larval instars, and after completion of larval development a pupal case is constructed, either by sealing off
the open end of the portable case (Integripalpia) or by
constructing a case and cementing it to the substrate
(Annulipalpia and Spicipalpia). The larva then ceases
feeding and becomes quiescent with its appendages
pressed against its body. This is the prepupal stage, and
may last a few days to several weeks. The pupa is of
the exarate type, with appendages free from the body.
Pupae in most families have well-developed mandibles
that the pharate adults (adult within the pupal integument) use to cut their way out of the case just prior to
emergence. After a pharate adult leaves the pupal case,
it swims to the water surface where eclosion takes
place or, more often, it swims to the surface and then
rapidly to shore where it crawls out of the water onto
some object to emerge. Eclosion is rapid.
Trichoptera larvae have many adaptations to their
aquatic environment. Respiration in caddisfly larvae is
through the integument, and in many families this is
aided by the presence of abdominal gills. Additionally,
especially in Integripalpia, larvae are able to increase
respiration by undulating their body within their tubular case to increase the flow of water and dissolved
oxygen through the case. Larval cases and retreats also
serve to protect larvae from predators, acting either as
a physical barrier or serving to camouflage the larva.
Cases may also protect larvae from their environment.
Hard cases protect those that live in sand and gravel
from abrasion, and streamlined cases or nets allow others to inhabit substrates in very swift currents. Netspinning larvae (Annulipalpia) use their nets to strain
algae and other food material from the current in
streams. Some caddisfly larvae (Leptoceridae) have
fringes of setae on their legs that allow them to swim.
Others (some Brachycentridae) have fringes of setae on
their legs to permit them to filter food particles from
the current.
Most caddisfly larvae can be readily collected from
lotic and lentic habitats with an aquatic net by using
methods already described. Debris in the pan should be
676
W. L. Hilsenhoff
examined carefully as many larvae have cases that blend
in with the debris. In streams it is important to also examine large substrates such as wood and rocks, because
larvae of many species attach themselves firmly to these
substrates or inhabit moss or algae that grows on them.
Species of larvae that burrow into decaying wood can be
collected as they crawl from retreats when the wood is
removed from a stream and allowed to dry. Larvae that
inhabit sand or silt substrates are best collected by sieving these substrates through a net or strainer, or by
closely watching sand substrates and collecting larvae
that expose themselves by moving. Larvae of extremely
small species are often overlooked because they remain
attached to the substrate and are too small to see. Microscopic examination of macrophytes and other substrates returned to the laboratory may yield many small
larvae, especially those of Hydroptilidae. In the field a
hand lens is helpful in locating very small specimens.
Caddisflies are a very important part of the stream
community; in some streams they dominate the insect
biomass. Many species of fish feed on the larvae and
emerging adults. Because of their general abundance
and diversity in streams, and the wide variation in tolerance to pollution among larvae of various species,
they are very important in biological monitoring. Occasionally, extremely large numbers of adults emerge, and
because they are attracted to lights they may create severe problems. These include contamination of products in factories, annoyance of people in restaurants
and elsewhere, and allergic reactions in people who are
sensitive to them.
Larvae of many species have never been associated
with adults. Larvae of these species must be reared,
associated with adults, and described before we will be
able to develop reliable keys to larvae at the species
level. Such keys are needed to enable us to complete
needed studies of the life history, ecology, and behavior
of the various species. Relatively little is presently
known about the life history and ecology of most North
American species. Most of what is known has resulted
from studies within a single stream where the fauna is
limited and thoroughly known because larvae have
been associated with adults. Wiggins (1996) summarized knowledge of the biology, ecology, morphology,
and distribution of larvae in each genus. Information on
the biology and habitat, and a brief description of the
larva in each caddisfly family follows. A key to families
of freshwater Trichoptera larvae appears in Section V.F,
and references to species keys for the identification of
larvae are provided in Table IV.
1. Families of Trichoptera — Suborder Annulipalpia
a. Dipseudopsidae (1 Genus, 5 Species) The only
genus Phylocentropus, was sometimes included in the
TABLE IV Literature References to Species Keys for
Identification of Trichoptera Larvae
Regional keys to larvae that include more than one family
Florida — Pescador et al. (1995)
Illinois — (many incomplete) Ross (1944)
North and South Carolina — Unzicker et al. (1982)
Keys to larvae listed by family
Apataniidae
Apatania — Chen (1992)
Brachycentridae — (Wisconsin) Hilsenhoff (1985)
Brachycentrus — Flint (1984)
Micrasema — Chapin (1978)
Goeridae
Goera — (eastern United States) Flint (1960); Wiggins (1996)
Goerita — Parker (1998)
other genera — Wiggins (1973)
Hydropsychidae — (Wisconsin) Schmude and Hilsenhoff (1986)
Arctopsychinae — Flint (1961); (western North America)
Givens and Smith (1980)
Ceratopsyche (as Hydropsyche) — Schefter and Wiggins (1986)
Hydropsyche (including Ceratopsyche) — (eastern and central
North America) Schuster and Etnier (1978)
Lepidostomatidae — Weaver (1988)
Leptoceridae
Ceraclea — Resh (1976)
Mystacides — Yamamoto and Wiggins (1964)
Nectopsyche — Haddock (1977)
Oecetis — Floyd (1995)
Setodes — Nations (1994)
Triaenodes and Ylodes — Glover (1996)
Limnephilidae — (eastern United States) Flint (1960)
Desmona — Wiggins and Wisseman (1990)
Dicosmoecus — Wiggins and Richardson (1982)
Hesperophylax — Parker and Wiggins (1985)
Pycnopshche — Wojtowicz (1982)
Molannidae
Molanna — Sherberger and Wallace (1971)
Odontoceridae
Psilotreta — Parker and Wiggins (1987)
Philopotamidae — Ross (1944)
Phryganeidae — Wiggins (1960)
Polycentropodidae
Paranyctiophylax — (as Nyctiophylax) Flint (1964);
Wiggins (1996)
Psychomyiidae
Psychomyia — Flint (1964)
Rhyacophilidae — (eastern United States) Flint (1962);
(Pennsylvania) Weaver and Sykora (1979)
generally predaceous family Polycentropodidae. Larvae
differ from those of Polycentropodidae by having
broad, flattened, setose tarsi (Fig. 72) and short, broad
mandibles with mesal brushes. These are adaptations
for their feeding on detritus, which they collect with a
net placed within tubes that they construct in sand and
silt along margins of sandy streams and lakes. The
capture net and feeding habits were described by
Wallace et al. (1976). All North American species
17. Diversity and Classification of Insects and Collembola
occur east of the Great Plains. The life cycle is probably
1 year, with cohorts emerging from May through
August.
b. Ecnomidae (1 Genus, 2 Species) Larvae of this
Neotropical family were collected from a small, cold,
rapid stream in the “hills” section of Texas (Waltz and
McCafferty, 1983). A second species was described by
Bowles (1995). These represent the only records of this
family north of Mexico. While previously considered a
subfamily of Polycentropodidae, larvae have sclerotized
meso- and metanota as do larvae of Hydroptilidae and
Hydropsychidae.
c. Hydropsychidae (11 Genera, 158 Species) Larvae
in this very large and important family are easily recognized by the numerous, branched filamentous gills that
occur ventrally on their abdomen. They frequently
represent a large segment of the macroinvertebrate
fauna in streams of all sizes, currents, and temperatures. Here, they build retreats on rocks and other
larger substrates, in leafpacks, or in moss. Larvae of
some species also tunnel into decaying wood and others may occur on wave-swept lake shores. Larvae of
most species are omnivorous, feeding primarily on algae, crustacea, and insects collected by their capture
nets; those of a few species tend to eat mostly algae and
diatoms and others are mostly predaceous, especially in
later instars. Life cycles vary from bivoltine to semivoltine, depending very much on stream temperature, with
the majority of species probably being univoltine. Because of their widespread abundance and the great
variation in the tolerance of species to organic pollution, larvae are very important in biological monitoring
of streams.
The subfamily Arctopsychinae is generally recognized in Europe as a separate family, and in North
America this family has been recognized by Schmid
(1968) and Nimmo (1987), but Wiggins (1981) presented compelling arguments for retention of subfamily
status. Capture nets of Arctopsychinae larvae are generally larger than those of other Hydropsychidae and
larvae are predominantly predators in the later instars.
Larvae differ from those of other Hydropsychidae by
having an elongate, posteriorly narrowed gula that
completely separates the genae.
d. Philopotamidae (3 Genera, 42 Species) Larvae
live in silken retreats, usually on the bottom of rocks in
warm-to-cold streams of all sizes. Their unmarked
head with an unsclerotized T-shaped labrum is distinctive (Fig. 70) and readily separates them from other net
spinning larvae with unsclerotized meso- and metanota. Larvae feed on algae and detritus that they capture
677
in their silken nets, which have a smaller mesh size
than those of other net spinning caddisflies. Most, if
not all species, are probably univoltine, except in the
south, where two or more generations may occur. Cohorts of most species emerge throughout the spring and
summer months; one species also emerges in fall and
winter.
e. Polycentropodidae (6 Genera, 71 Species) While
larvae of most species live in streams, they inhabit a
wide variety of other habitats, including littoral areas
of lakes and temporary ponds. Larvae can be recognized by their pointed protrochantin (Fig. 71), unmodified tarsi, and the presence of dark or light spots (muscle scars) on the heads of most species. Reliable keys to
larvae do not exist, because larvae of many species remain undescribed. Most larvae have a waxy cuticle
that tends to repel water, giving the impression when
collected that they are terrestrial. Larvae of a few
species build trumpet-shaped capture nets similar to
other Annulipalpia and are primarily herbivores, but
those of most species are predators and have tubular
retreats that they sometimes use to detect prey. These
tubular retreats also function as an aid to respiration,
with larvae circulating water through them by undulating their body. This is especially important for species
that live in lentic habitats that occasionally may be
deficient in oxygen. Life cycles are probably one year in
most species, with emergence during the summer
months.
f. Psychomyiidae (4 Genera, 17 Species) Larvae
live in streams, frequently on or in wood; they also occur on rocks, which they cover with silken retreats that
incorporate sand or detritus. Larvae can be readily separated from those of other families without sclerites on
the meso- and metanotum by their hatchet-shaped protrochantin (Fig. 69). They graze food from the vicinity
of their retreats, mostly consuming detritus, fungi, and
periphyton. Most species are probably univoltine, with
emergence in late spring and summer.
g. Xiphocentronidae (2 Genera, 2 Species) Sometimes considered as part of Psychomyiidae, the only described larvae of Xiphocentronidae north of Mexico
differ from Psychomyiidae larvae and those in other
families by having their tarsi and tibiae fused into a single segment on all legs (Fig. 68). Larvae are known in
North America from southern Texas, where one species
was collected from a spring run and described by
Edwards (1961), and from northern Arizona, where
larvae in a second genus were described by Moulton
and Stewart (1997).
678
W. L. Hilsenhoff
2. Families of Trichoptera — Suborder Integripalpia
a. Apataniidae (5 Genera, 33 Species) Until recently this family was included in Limnephilidae; the
larvae are quite diverse morphologically. All are rather
small (12 mm) and live in cornucopia-shaped cases of
small rock fragments. They occur in small cold-water
streams or seeps, and occasionally in cold lakes where
they feed mostly on algae. Most larvae differ from
other Limnephilidae by having mandibles with uniform
scraper blades; those with toothed mandibles have 25
or more setae on the anterior of the metanotum between the sclerites.
b. Beraeidae (1 Genus, 3 Species) The gently
curved and tapered case is made of fine sand. Larvae
are small and can be recognized by a transverse ridge
on the pronotum that curves forward laterally to form
rounded lobes (Fig. 77). Larvae, which are known for
only two North American species (Wiggins, 1996), inhabit muck margins of spring seeps. Adults have been
collected only in southeastern and northeastern North
America, where they are rare. A species found in Ontario has been reported to be semivoltine.
c. Brachycentridae (5 Genera, 34 Species) The larval case is tapered, of vegetable matter or sand grains,
and square or round in cross section. Larvae are distinctive, having their pronotum divided by a distinct
furrow (Fig. 76), with the area in front of the furrow
depressed. They have no dorsal or lateral humps on the
first abdominal segment, which is unique among Integripalpia. Larvae in three of the genera are monotypic
in North America; most others can be identified by
keys listed in Table IV. Most species are univoltine, but
some that inhabit cold northern streams are semivoltine. Larvae of all species inhabit streams, especially
cold spring-fed streams, where they live in moss or
other vegetation, or attach their cases to substrates in
the current. The mostly herbivorous larvae graze on periphyton and other vegetation; some also consume
small insects and crustaceans. Some are primarily filter
feeders that use their elongate meso- and metathoracic
legs to gather food from the current.
d. Calamoceratidae (3 Genera, 5 Species) Cases are
of leaf pieces or hollowed sticks. Larvae, which may
reach a relatively large size, can be recognized by the
projecting, divergent, anterolateral corners of the
pronotum and a row of about 16 long setae dorsally
on the labrum (Fig. 79). Most species can be identified
as larvae by using descriptions and distributions reported in Wiggins (1996). Larvae inhabit pool areas
of cool streams in the west or east, where they feed
mostly on detritus. Most species probably have a
2-year life cycle.
e. Goeridae (4 Genera, 11 Species) Larvae resemble those of Limnephilidae, but differ in their unique
modification of the mesepisternum, which is formed
into an anteriorly projecting elongate process (Fig. 83)
or a rounded spiny prominece. Since there are only a
limited number of species, it is often possible to
identify larvae to species using publications listed in
Table IV. The sand cases are either cornucopia-shaped
or tubular with flanges of pebbles. Life cycles are probably univoltine or semivoltine. Larvae inhabit cobbles
and other substrates in the current of small cold
streams; larvae of one monotypic genus occur in the
muck of spring seeps. They feed on algae, vascular
plants, or detritus.
f. Helicopsychidae (1 Genus, 4 Species) The snail
like case, which larvae never abandon, is unique (Fig.
59). Only the larva of Helicopsyche borealis, which is
widespread in North America, is known; larvae of the
three western species remain unknown. The normally
lotic larvae may also be found along margins of lakes,
often in rather deep water. Larvae inhabit rocks or
wood and scrape periphyton and detritus from the substrate; early instars may burrow into the bed of sandy
streams. They can tolerate very warm water, and have
been collected from thermal springs with temperatures
as high as 34°C. They are probably univoltine, with
emergence of cohorts throughout the summer and a
prolonged winter diapause of eggs in the North.
g. Lepidostomatidae (2 Genera, 82 Species) Cases
are highly variable, being made of sand or bits of vegetation and square or round in cross section. The location of the antenna very close to the eye (Fig. 78) readily identifies all larvae. Larvae are primarily detritivores
and are found among detritus on a variety of substrates. While larvae of most species inhabit cold
streams or springs, some occur in lakes and may be
found at a considerable distance from the shore.
Species that have been studied are univoltine, with spatial or temporal separation of different species that occur in the same stream.
h. Leptoceridae (8 Genera, 113 Species) Larvae
construct a wide variety of cases. In some genera they
have elongate cases of vegetation, vegetation and sand,
or of pure silk, and are adapted for swimming by having fringes of long setae on their legs. Larvae in other
genera have shorter, stouter cases that are often cornucopia-shaped and constructed of vegetable or mineral
material; these larvae are unable to swim. Most larvae
17. Diversity and Classification of Insects and Collembola
can be recognized by their relatively long antennae
(Fig. 63), which are much longer than those of other
caddisfly larvae, and also by their elongate metathoracic legs, which are distinctly longer than their other
legs. Larvae inhabit a variety of permanent lotic and
lentic habitats; those in lotic habitats tend to be in
slower water and those in lakes may be found far from
shore as well as in the littoral zone. They are generally
omnivorous in their feeding habits, but some are primarily predators and a few feed exclusively on freshwater sponges. Most species are probably univoltine.
i. Limnephilidae (39 Genera, 243 Species) Case
construction in this large and diverse family varies
widely, with vegetable or mineral material incorporated
into a variety of shapes and sizes. Larvae also vary
greatly in size and structure, making them difficult to
characterize. All have a prosternal horn and antennae
that are located about halfway between each eye and
mandible (Fig. 80). Most have numerous setae on the
first abdominal segment, and sclerites at SA-1 of the
metanotum (Fig. 66). Many also have chloride epithelia
ventrally or dorsally on abdominal segments. While
species in some genera can be identified by keys listed
below, accurate identification of larvae is not possible
in most genera because larvae of too many species remain unknown. Most species are univoltine, but a few
are semivoltine. Larvae of most species inhabit a wide
range of lotic habitats, from springs and small streams
to large rivers; others inhabit lakes, and several are
found in permanent or temporary ponds. In these habitats larvae can be found on vegetation, rocks, or detritus, or in gravel, sand, or soft sediments. Most larvae
are herbivores or detritivores, and are important in
processing large particulate organic matter; larvae of
one species are known to be carnivores.
j. Molannidae (2 Genera, 7 Species) Larvae construct characteristic sand cases with lateral flanges and
a hood over the anterior opening. They also have long
metathoracic legs with either very short or threadlike
claws. Larvae live in sandy or silty substrates of lakes
or streams where currents are reduced. Here they feed
on algae, vascular plants, or even invertebrates. Most
species are probably univoltine.
k. Odontoceridae (6 Genera, 13 Species) Cases are
typically cornucopia-shaped and made of sand grains
and tiny stones; they are hard and difficult to crush.
Like Sericostomatidae, which they resemble, larvae
lack a prosternal horn and have their eyes located close
to the mandibles, but their protrochantin is small and
not hook-shaped (Fig. 82). Species of larvae in the
largest genus (Psilotreta) can be identified (Table IV);
679
species in monotypic genera can be identified by using
the key preposed by Wiggins (1996). Larvae inhabit
small, cold streams or springs where they are found in
sandy substrates. They are probably omnivorous, feeding on algae, vascular plants, and invertebrates. Most
species are probably semivoltine in the north and univoltine in the south.
l. Phryganeidae (10 Genera, 28 Species) These
large caddisflies have distinctive cases made mostly of
pieces of vegetation that are spirally wound or in concentric rings. When disturbed, they readily abandon
their cases, but may re-enter them. Larvae are distinctive, generally having a boldly striped head that is more
prognathous than other Integripalpia (Fig. 65). They
also have a prominent prosternal horn and lack significant sclerotization of the mesonotum. Identification of
larvae to species is not possible in some genera. Larvae
may be found among vegetation and detritus along
streams of all sizes, in marshes, in temporary and permanent ponds, and even in lakes where they may occur
far from shore. While many are mostly predators, vegetation is also consumed, especially by early instars. Life
cycles are probably 1 year.
m. Rossianidae (2 Genera, 2 Species) This western
North American family was erected for two genera previously placed in Limnephilidae and Goeridae. Larvae
live in organic muck of spring seeps or in small, cold,
mountain streams where they feed on plants and detritus. Their curved cases are made of tiny stones. Larvae
are distinctive among Integripalpia because their head,
pronotum, and mesepisternum are modified by prominent surface sculpturing.
n. Sericostomatidae (3 Genera, 15 Species) Cases
are typically cornucopia-shaped and made of sand and
small pebbles, but are not hard as in Odontoceridae.
Larvae can be recognized by the pointed anterolateral
angles of their pronotum, a hook-shaped protrochantin
(Fig. 81), and broad, flat profemur. No key has been
published for separating larvae of Agarodes, the largest
genus. Larvae live in sandy substrates of streams,
springs, and even lake margins, where they feed primarily on plant detritus. Most species are probably
univoltine.
o. Uenoidae (5 Genera, 46 Species) The cornucopiashaped cases of sand grains are either extremely long and
thin or short and stout, depending on the genus. Larvae
occur in streams and springs. Until recently they were
considered part of Limnephilidae. Larvae differ from
Limnephilidae by having a mesally notched anterior of
the mesonotum (Fig. 84) and untoothed scraping
680
W. L. Hilsenhoff
mandibles, which they use to scrape diatoms and organic
detritus from rock surfaces. There are several species in
each of the genera, and keys to species of larvae have not
yet been developed. Two genera (Neothremma and
Farula) occur in the western mountains; Neophylax, a
species widespread in North America, was recently
added to this family (Vineyard and Wiggins, 1988).
ognized by their unsclerotized meso- and metanotum,
sclerotized ninth abdominal tergum, and elongate anal
prolegs (Fig. 74). Life cycles are probably univoltine for
most species, but may be semivoltine in some. Larvae
of most species are active predators, but those of a few
species feed on living and dead vascular plant tissue
and algae.
3. Families of Trichoptera — Suborder Spicipalpia
a. Glossosomatidae (6 Genera, 76 Species) Larvae
build unique turtlelike cases of sand and pebbles, but
readily vacate them when collected. They can be separated from other Spicipalpia with unsclerotized mesoand metanota by their cases and their short anal prolegs with short claws (Fig. 73). Larvae can be identified
to species only by associating them with adults. All
species inhabit the upper surfaces of rocks or other
large substrates in clean, cool streams or springs. Here
they feed on diatoms and other algae that they scrape
from the substrate. Most species probably have a oneyear life cycle, often with many cohorts that emerge
throughout the spring and summer.
b. Hydrobiosidae (1 Genus, 3 Species) Although
sometimes included in Rhyacophilidae, the free-living
larvae are unique in having chelate prothoracic legs
(Fig. 75), which are suited to their predatory habits.
They occur in cool streams in Texas, Arizona, and
Nevada. The larva of one species was described by
Edwards and Arnold (1961).
c. Hydroptilidae (16 Genera, 250 Species) Larvae
are characterized by their completely sclerotized thoracic nota, lack of branched ventral gills, and very
small size. They are frequently called “microcaddisflies.” The first four larval instars are free-living, have
no case, and have a normal-size abdomen; in the last
instar the abdomen is greatly expanded and larvae have
a case that is purse-shaped or barrel-shaped and sometimes attached to the substrate (Figs. 61, 62). Most
species pass through the first four instars rapidly, and it
is usually not possible to identify these early instars to
genus. Larvae in the last instar can be identified to
genus, but usually not to species. Larvae may be found
in a wide variety of lotic and lentic habitats where they
feed mostly on algae and other plant material. They are
probably univoltine, or bivoltine in warmer habitats,
but generally very little is known about life histories of
the various species.
d. Rhyacophilidae (2 Genera, 128 Species) The larvae live in cool or cold streams, have no case, and do
not build retreats. Most species occur in mountainous
regions where cold, fast streams abound. They are rec-
E. Megaloptera — Fishflies and Alderflies
This very small holometabolous order contains
two aquatic families that once were included in the order Neuroptera. The aquatic larvae are distinctive, having seven or eight pairs of lateral filaments and large,
conspicuous mandibles (Figs. 85, 86). They can be confused only with some aquatic Coleoptera larvae that
have lateral filaments. Larvae often attain a very large
size, and because all are predators, they may exert significant pressure on populations of macroinvertebrates
that live in the same habitat. Studies indicate that there
are usually 10 or 11 larval instars, and that life cycles
range from 1 – 4 years, depending on the species, climate, and habitat. Larvae are aquatic; all other stages
are terrestrial. Eggs are laid in masses on objects above
the water, and pupation takes place in cells on land.
The terrestrial adults are short-lived and generally secretive, but because of their large size and often conspicuous mandibles, they attract attention when discovered. Adults are weak fliers, but are often attracted to
lights. They fold their net-veined wings rooflike over
their abdomen when at rest.
Larvae of most species occur only in relatively oxygen-rich environments, because they usually obtain
oxygen through their integument directly from the water in which they live. Their lateral filaments greatly increase surface area, allowing them to do this. Species
that inhabit lentic habitats, which may at times be low
in oxygen, have long caudal respiratory tubes that they
use to obtain oxygen from air at the surface. All larvae
have spiracles and are capable of living out of water in
moist areas; larvae of some species survive in temporary streams.
Larvae can be readily collected with an aquatic net
by using standard sampling procedures. However,
greater numbers of stream species can often be collected by dislodging larger rocks. Rearing of larvae is
difficult because of long life cycles, cannibalism, and
the need to supply prey species and a terrestrial environment for pupation. Species of adults are well
known, and as a result of recent studies larvae of most
species can also be identified. References to species keys
for the identification of larvae are provided in Table V.
Information on the biology, habitat, and identification
of larvae in the two families is presented next, and a
17. Diversity and Classification of Insects and Collembola
TABLE V Literature References to Species Keys
for Identification of Megaloptera Larvae
Regional keys to larvae that include more than one family
North and South Carolina — Brigham (1982a)
Keys to larvae listed by family
Corydalidae — (Mississippi) Stark and Lago (1983)
Chauliodes — Cuyler (1958)
Nigronia — Neunzig (1966)
Sialidae — (eastern North America) Canterbury (1978)
key to families of freshwater Megaloptera larvae appears in Section V.F.
1. Families of Megaloptera
a. Corydalidae (7 Genera, 22 Species) The very
large size (20 – 90 mm), lateral filaments, and large
mandibles of late instar “fishfly” or “Dobsonfly” larvae
are distinctive (Fig. 86), but smaller larvae resemble
those of Gyrinidae and two other Coleoptera genera.
Larvae can be distinguished from Coleoptera larvae by
their pair of terminal prolegs, each with two hooks (Fig.
86). Larvae in most genera inhabit streams or spring
seeps, which have a relatively high dissolved oxygen
content. Because these habitats are generally cold, life
cycles of two or more years are likely for most species,
with different cohorts and extended emergence periods
in some species. The habitat of Chauliodes larvae differs
from that of other genera. They inhabit shallow lentic
habitats or vegetated margins of streams and are univoltine, with emergence in late spring or early summer.
They also are known to consume significant amounts of
detritus in addition to prey, especially during the winter.
b. Sialidae (1 Genus, 24 Species) The lateral filaments and long, single tail filament at once separate
“alderfly” larvae from all other aquatic insects (Fig.
85). Larvae live mostly in depositional sediments of
streams, permanent ponds, and lakes. Species that live
in lakes may inhabit areas that are several hundred meters from shore, necessitating a considerable migration
to land for pupation. Most species are univoltine, but
in northern regions species that live in streams or lakes
are often semivoltine.
III. PARTIALLY AQUATIC ORDERS OF INSECTS
A. Aquatic Heteroptera — Aquatic
and Semiaquatic Bugs
Because all aquatic and semiaquatic bugs are Heteroptera and none are Homoptera, the catalog by
Henry and Froeschner (1988) will be followed and this
681
order of insects will be referred to as Heteroptera
rather than Hemiptera, or Hemiptera: suborder Heteroptera. The choice of names for this group of insects
has created much controversy; pros and cons are discussed by Henry and Froeschner (1988).
Heteroptera is a medium-sized paurometabolous
order that is primarily terrestrial, but about 8.5% of
the species are aquatic, with adults and nymphs either
living in water (220 species) or on its surface (106
species). It is divided into seven suborders, only two of
which have aquatic species. Species with adults and
nymphs that walk on the water surface are in the suborder Gerromorpha, and are referred to as “semiaquatic Heteroptera.” Species with adults and nymphs
that live in the water belong to the suborder Nepomorpha. These are referred to as “aquatic Heteroptera,”
because they spend their entire life under water, except
during dispersal flights. Flightless species of Nepomorpha are the most aquatic of all insects because they
rarely, if ever, leave the water. There are six families of
Nepomorpha in which all species are aquatic, and five
families of Gerromorpha in which all species are
semiaquatic or occasionally riparian. There are also
four families of Heteroptera with riparian adults
and nymphs; these have been included in the key
because they occasionally are collected along with
aquatic species. These are Gelastocoridae and Ochteridae (Nepomorpha), Macroveliidae (Gerromorpha), and
Saldidae (Leptopodomorpha).
Most species of aquatic and semiaquatic Heteroptera breed in lentic habitats, but there are also several lotic species, and adults of many lentic species fly
to streams in the north to overwinter. Adults and
nymphs of most species are predators, and their presence can significantly influence populations of other insects, especially in lentic habitats. They are known to
have an impact on populations of mosquito larvae, and
thus can benefit man. Adults of larger species, however,
may prey on small fish and create problems in fish-rearing ponds. Those of some species also may bite people,
and occasionally they become a nuisance when attracted by lights to swimming pools. The vast numbers
of corixid adults that overwinter in northern streams
are an important source of food for some fish, and
dried corixids are sold as food for pet turtles and tropical fish. Most other Heteroptera are usually avoided by
fish, probably because of secretions from their scent
glands. Generally, however, aquatic and semiaquatic
Heteroptera have little impact on humans.
Heteroptera adults can be distinguished from other
aquatic insects by their mesothoracic wings, or hemelytra, which are hardened in the basal half and membranous apically (Fig. 87), and by sucking mouthparts that
are formed into a tube or “rostrum” (Figs. 88, 90).
682
W. L. Hilsenhoff
There are normally five nymphal instars; a few species
have only four. Nymphs resemble adults, except that
they lack wings and genitalia, and are distinctly less
sclerotized. They inhabit the same habitat as adults and
are often associated with them.
Aquatic and semiaquatic Heteroptera have been
rather thoroughly studied, and adults of all North
American species can be readily identified. Families and
genera are quite heterogeneous, making it possible to
recognize families, many genera, and some species in the
field. The nymphs, however, have not been well-studied
except for Gerridae, and few can be identified to species
except regionally or through association with adults.
Some cannot even be identified to genus. Since the pioneering works of H.B. Hungerford and others 45 – 80
years ago, there have been relatively few additions to the
known fauna in North America. References to species
keys to adults are provided in Table VI. Information on
the biology, habitat, and identification of each family is
presented next, along with information about suborders.
A key to families of adults of aquatic, semiaquatic, and
riparian Heteroptera appears in Section V.G.
1. Families of Semiaquatic Heteroptera — Suborder
Gerromorpha
Adults of semiaquatic species reach peak abundance in September or October, depending on latitude,
after which most of them find protected terrestrial sites
in which to spend the winter; adults of species that
overwinter as eggs die. Species that overwinter as
adults are bivoltine or multivoltine; those that overwinter as eggs usually are univoltine, but may be bivoltine
in the south. Eggs of semiaquatic species are laid on
substrates at the water surface. Nymphs and adults
feed mostly on invertebrates of an appropriate size in
the surface film. Gerridae, and probably most other
Gerromorpha, have sensillae on their femurs and
trochanters that detect vibrations in the surface film,
enabling them to find their prey. In most semiaquatic
Heteroptera apterous and brachypterous adults are
common. Apterous adults can be distinguished from
nymphs by the presence of genitalia, greater sclerotization of the body, and by having two tarsal segments instead of one on most legs.
Semiaquatic Heteroptera adults and nymphs are
adapted for walking on water by having hydrofuge
hairs on their tarsi, and in the larger and heavier Gerridae and Veliidae, by additionally having preapical
claws. They are most readily collected by sighting them
on the water surface and sweeping them from the surface film with an aquatic net.
a. Gerridae (8 Genera, 44 Species) All of the largest
adults of semiaquatic Heteroptera are species of Gerri-
TABLE VI Literature References to Species Keys
for Identification of Heteroptera Adults
Regional keys to adults that include more than one family
Alberta, Saskatchewan, and Manitoba — Brooks and
Kelton (1967)
California — Menke (1979a)
Florida — (northern) Herring (1951); Chapman (1958)
Minnesota — Bennett and Cook (1981)
Mississippi — Wilson (1958)
Missouri — Froeschner (1949, 1962)
Montana — Roemhild (1976)
North and South Carolina — Sanderson (1982)
Oklahoma — Schaefer and Drew (1968)
Virginia — Bobb (1974)
Wisconsin — Hilsenhoff (1984b, 1986)
Keys to adults listed by family
Belostomatidae – Menke (1979b); (Louisiana) Gonsoulin (1973c)
Corixidae – Hungerford (1948); (Oregon and Washington)
Stonedahl and Lattin (1986)
Cenocorixa – Jansson (1972)
Hesperocorixa – Dunn (1979)
Trichocorixa – Sailer (1948)
Gerridae – (Arkansas) Kittle (1980); (British Columbia) Scudder
(1971); (Louisiana) Gonsoulin (1974); (Oregon and
Washington) Stonedahl and Lattin (1982); (Ontario) Cheng
and Fernando (1970)
Aquarius – Andersen (1990)
Gerrinae – Drake and Harris (1934); Kuitert (1942)
Gerris – Andersen (1993); (Connecticut — also nymphs)
Calabrese (1974)
Limnoporus – Andersen and Spence (1992)
Metrobates – Anderson (1932)
Rheumatobates – Hungerford (1954)
Trepobates – Kittle (1977)
Hebridae – Porter (1950)
Hydrometridae – Hungerford and Evans (1934); (Louisiana)
Gonsoulin (1973a)
Mesoveliidae – Polhemus and Chapman (1979)
Naucoridae – (Texas) Davis (1996); (Louisiana) Gonsoulin (1973b)
Ambrysus – LaRivers (1951)
Pelocoris – LaRivers (1948); (Texas) Sites and Polhemus (1995)
Nepidae – Sites and Polhemus (1994); (Louisiana) Gonsoulin (1975)
Notonectidae – (Arizona) Zalom (1977); (British Columbia)
Scudder (1965); (West Coast nymphs) Voigt and Garcia (1976)
Buenoa – Truxal (1953)
Notonecta – Hungerford (1933); (New England)
Hutchinson (1945)
Veliidae – Smith and Polhemus (1978); Smith (1980)
Rhagovelia – Polhemus (1997)
dae, but the family has many diverse sizes, forms, and
life histories. Adult “water-striders” are distinguished
by their relatively large size, preapical tarsal claws (Fig.
97), and elongate metafemurs that extend well past the
tip of the short abdomen. Apterous and brachypterous
forms are common. Although found in all types of
aquatic habitats, adults and nymphs of most species
prefer quiet areas where there is minimal wave action
and current. Adults and nymphs of lotic species inhabit
17. Diversity and Classification of Insects and Collembola
streams of all sizes; those of lentic species inhabit
swamps, marshes, permanent and temporary ponds, littoral areas of lakes, and even the ocean. However,
adults of most lotic species will occasionally inhabit
lentic habitats and adults of lentic species may occasionally be found along margins of lotic habitats. Most are
usually associated with emergent vegetation. Univoltine
or bivoltine life cycles prevail, with some species overwintering as adults and some univoltine species overwintering as eggs. All water striders are predaceous.
They feed on terrestrial insects and other invertebrates
that become trapped in the surface film or aquatic insects that live just beneath the surface.
b. Hebridae (3 Genera, 15 Species) Because of
their very small size (2.5 mm), “velvet water bugs”
are likely to be confused only with very small species of
Veliidae. When on the water, Hebridae adults and
nymphs move much more slowly than those of similarsized Veliidae, and upon close examination Hebridae
adults and nymphs have a rostral sulcus and apical
claws, which those of Veliidae lack. Adults and nymphs
inhabit very shallow, sheltered shoreline areas of lentic
habitats, including swamps, marshes, ponds, lakes, and
river sloughs. Those of some species are occasionally or
even frequently riparian. Apterous adults are common
in some species. Adults can be found from spring to autumn, suggesting that they are bivoltine or multivoltine
and overwinter as adults.
c. Hydrometridae (1 Genus, 9 Species) The small
(8 – 11 mm), delicate, sticklike “marsh-treaders” or
“water-measurers” with their elongate head (Fig. 99)
are unlike any other aquatic insect. Both macropterous
and brachypterous adults are common. They walk on
vegetation and the surface film of permanent habitats
that lack wave action and current, including marshes,
swamps, and vegetated margins of lakes, ponds, and
slow streams. Here they feed by using their barbed rostrum to spear small macroinvertebrates in the surface
film. They most often are found where fish are absent.
Hydrometridae are multivoltine and overwinter as
adults.
d. Mesoveliidae (1 Genus, 3 Species) “Water-treaders” become abundant in late summer in sheltered
lentic habitats and margins of lotic habitats that have
algae and duckweed along with emergent vegetation.
Their relatively small size (2 – 4 mm) and the presence
of black spines on the legs is distinctive. Apterous
adults predominate, but winged adults are not uncommon, especially in autumn. They prey on small terrestrial and aquatic animals in the surface film. Adults and
nymphs are killed by freezing temperatures, and in the
683
north mesoveliids overwinter as eggs. All species are
probably multivoltine.
e. Veliidae (5 Genera, 35 Species) “Broad-shouldered” or “short-legged water-striders” are often abundant in a wide variety of aquatic habitats. Some inhabit
streams, living in the vicinity of riffles or under the
banks; others inhabit spring ponds, swamps, marshes,
ponds, and margins of lakes. Adults can be distinguished from other semiaquatic Heteroptera, except
Gerridae, by their preapical tarsal claws. They differ
from Gerridae in their generally smaller size and relatively shorter legs, the metafemurs not extending past
the tip of the abdomen. Apterous adults predominate,
with winged forms rare or absent in the various
species. Most species are multivoltine and overwinter
as adults, some along margins of streams or springs instead of in terrestrial habitals. A few species overwinter
as eggs and are univoltine or bivoltine.
2. Families of Aquatic Heteroptera — Suborder
Nepomorpha
Most species are univoltine in northern North
America; a few are bivoltine. Farther south, bivoltine
life cycles probably predominate. Adults reach peak
abundance in October or November, and overwinter in
aquatic habitats. In areas where ponds and lakes freeze,
adults of species that typically breed in shallow lentic
habitats usually fly to deep lentic habitats or larger
streams to overwinter. In late April or early May, they
return to their breeding sites to mate and oviposit. Eggs
are laid on vegetation or other substrates, usually in the
water. Some univoltine species diapause over the winter
as eggs.
Adults and nymphs of most species are adapted for
swimming by having fringes of long setae on their legs,
especially the metathoracic legs. They obtain oxygen
from air stores held under the hemelytra and from a
bubble held ventrally on hydrofuge hairs, which may
act as a physical gill or plastron. Air stores are renewed
at the surface through air straps in Belostomatidae, air
tubes in Nepidae, the pronotum in Corixidae, and the
apex of the abdomen in Notonectidae and Naucoridae.
In Corixidae, Notonectidae, Pleidae, and Naucoridae
the ventral bubble is large in relation to their size and
acts as a physical gill, with oxygen from the water diffusing into the bubble and carbon dioxide from respiration diffusing out of it. This allows prolonged submergence in oxygen-rich water. Adults and nymphs of
almost all species are carnivores. They feed mostly on
small invertebrates and occasionally on vertebrates
such as minnows and tadpoles. The largest family,
Corixidae, is an exception, with adults and nymphs of
most species being primarily detritivores.
684
W. L. Hilsenhoff
Nepomorpha adults and nymphs are best collected
with an aquatic net along with debris. The contents of
the net are then spread on a half-inch (13 mm) mesh
screen over a large white pan, and insects are allowed
to crawl from the debris and fall into the pan. Adults
and nymphs of many species feign death and will crawl
from the debris only if left undisturbed for several minutes. Additional specimens in all families can often be
collected with bottle traps (Hilsenhoff, 1987). In northern regions, exceptionally large numbers of adults of
both lotic and lentic species can be collected in autumn
from along banks of large streams where the current is
slow, but adults of some lentic species do not fly to
streams and will be missed. Adults of many species are
attracted to lights, and may be collected with light
traps.
permanent ponds and littoral areas of small lakes, but
those of some species are strictly lotic and can be found
only along margins of streams or in spring ponds and
seeps. In northern latitudes, adults of most lentic
species fly to larger streams to overwinter, and in
October and November thousands can often be collected from slow, shallow areas of streams. Adults disperse widely, rapidly invading temporary ponds, and
on warm nights they are readily attracted to lights.
Eggs are laid on underwater structures in such great
numbers that they are harvested for food in some tropical countries. Most northern species are probably univoltine, with overwintering adults returning to breeding sites in spring to mate and oviposit. In the south,
two or more generations per year are likely. There is
evidence that at least one species overwinters as an egg.
a. Belostomatidae (3 Genera, 19 Species) “Giant
water bugs” are easily recognized by their large size
and a pair of short, straplike, posterior respiratory appendages (Fig. 92) that they use to obtain air at the water surface. Adults and nymphs of two genera (Belostoma and Lethocerus) inhabit permanent lentic
habitats, especially weedy ponds, margins of lakes, and
marshes, and are important as apex invertebrate predators. Those of the third genus (Abedus) occur in
streams among aquatic plants or under rocks in riffles.
All adults are powerful predators that will capture and
kill anything they can subdue, including fish, frogs,
tadpoles, and other insects. Eggs are laid in large
masses above the water on vegetation (Lethocerus) or
on the backs of the males, which brood the eggs
(Abedus and Belostoma). Belostomatids are univoltine
or bivoltine, depending on the latitude, with adults of
lentic species in northern regions flying in autumn to
larger streams or deep lentic habitats where they overwinter. They return to their breeding sites in early May.
Adults frequently fly to lights and are sometimes called
“electric light bugs.”
c. Naucoridae (4 Genera, 22 Species) “Creeping
water bugs” are most common in the southern United
States and are rarely found as far north as Canada.
They are dorsoventrally flattened, have greatly expanded profemurs, and have a rounded appearance
when viewed from above, with margins of the head,
pronotum and elytra being continuous (Fig. 93). They
are usually associated with lotic habitats, living in
streams, spring ponds, or impoundments. Eggs are
glued to vegetation or mineral substrates, depending on
the species. Most species probably are univoltine
and overwinter as adults in deeper areas of their breeding habitat. Because adults carry a large plastron and
generally live in well-oxygenated water, they rarely
need to surface for air. Although adults of many species
have fully developed wings, flight rarely has been
observed.
b. Corixidae (18 Genera, 129 Species) As the most
abundant aquatic Heteroptera, “water boatmen” are
found in most permanent aquatic habitats and frequently invade temporary ones as well. Adults most
closely resemble those of Notonectidae, but are
dorsoventrally flattened (Fig. 87) and have a short,
broad, unsegmented, triangular rostrum and widened,
scoop-shaped protarsi (palae) (Fig. 88). Unlike other
aquatic and semiaquatic Heteroptera, which are predators, corixid adults and nymphs feed primarily on detritus, algae, protozoans, and other extremely small animals. Adults of a few species, however, will capture
and eat larger insects, such as mosquito larvae. Adults
and nymphs of most species are lentic, inhabiting
d. Nepidae (3 Genera, 13 Species) “Water scorpions” can be readily recognized by their long apical
breathing tube (Fig. 91). They breed in permanent
lentic habitats, especially shallow areas with much vegetation. Here they often remain with their breathing
tube above the surface waiting to ambush invertebrates
or small vertebrates, but they are capable of remaining
completely submerged, using air stores under their
hemelytra for respiration. Unlike other aquatic Heteroptera, adults and nymphs are very poor swimmers
and usually cling to vegetation or other substrates.
Most species are univoltine in northern latitudes and
probably bivoltine in the south. They overwinter as
adults, which in the north usually fly to deeper lentic or
lotic habitats in autumn and return to breeding sites in
late April or early May. Eggs that are laid on supports
at the water surface have respiratory horns that protrude from the water. When flooded, the eggs respire
through a plastron.
17. Diversity and Classification of Insects and Collembola
e. Notonectidae (3 Genera, 32 Species) “Backswimmers,” as their name implies, swim ventral side
up, but orient themselves dorsal side up when not in
the water. They most resemble Corixidae, but are deepbodied, not at all flattened (Fig. 95), and have an elongate, segmented rostrum (Fig. 90). Adults of most
species breed in littoral areas of lakes and permanent
ponds, others in stream pools; some species are transients in temporary ponds. Adults and nymphs of two
genera, Buenoa and Notonecta, can inhabit relatively
deep water and remain submerged for long periods because they have ventral channels to retain a plastron.
Notonecta adults and nymphs typically hang from the
surface film, while Buenoa adults and nymphs have hemoglobin, which enables them to maintain neutral
buoyancy and remain planktonic. Adults and nymphs
in both genera are active predators that pursue or ambush other invertebrates or small vertebrates. A third
genus, Martarega, occurs only in extreme western
United States, where adults and nymphs inhabit eddies
of streams and ambush prey carried by the current.
Several species overwinter as diapausing eggs and have
a univoltine life cycle with nymphal development in
late spring and summer. Others have univoltine or bivoltine life cycles, with overwintering adults that mate
and lay eggs in the spring. In the north, adults of a few
species may overwinter in larger streams and return to
lentic habitats in spring.
f. Pleidae (2 Genera, 5 Species) “Pygmy backswimmers” are very small (3 mm), convex insects
that swim upside down. Adults do not have typical
hemelytra, but instead their mesothoracic wings are
shelllike (Fig. 94) and resemble elytra of Coleoptera.
Their metathoracic wings are vestigial and they cannot
fly. Adults and nymphs inhabit vegetation, mostly in
permanent ponds, but also may be found in littoral areas of lakes, backwaters of streams, and swamps. If
their habitat dries because of drought, they can survive
for several months in the moist soil. Pleids are widespread and often abundant, except in the Pacific Coast
region where they do not occur. Although adults and
nymphs can swim quite well, they mostly crawl about
on vegetation and feed on small invertebrates. Because
of their small body size and relatively large plastron,
they are able to remain submerged for long periods.
Life cycles are bivoltine or multivoltine, but their life
history is poorly known.
B. Aquatic Neuroptera — Spongillaflies
Neuroptera is a fairly large terrestrial order, but
larvae in only one family are aquatic. They live in and
feed upon several genera of freshwater sponges
685
(Spongillidae) that occur in permanent lotic or lentic
habitats. Taxonomic keys to species of larvae can be
found in the following publications: Parfin and Gurney
(1956), Poirrier and Arceneaux (1972), and for North
and South Carolinas in Brigham (1982b). Information
on the biology, habitat, and identification of larvae is
presented next.
1. Families of Aquatic Neuroptera
a. Sisyridae (2 Genera, 6 Species) The small
(8 mm) aquatic larvae are distinctive. They have
sucking mouthparts in the form of a pair of long stylets
and are covered with long, spinose setae (Fig. 102).
Most species are probably multivoltine, with larvae
having only three instars. Other life stages are terrestrial. Eggs are usually laid in very small masses, mostly
in crevices of vegetation or supports above aquatic
habitats that support freshwater sponges. After an incubation period of about 1 week, the eggs hatch and
larvae drop into the water where they swim or are
carried by currents until they encounter a suitable
species of sponge. After completing development on the
sponge, they swim or crawl to shore, crawl out of
the water, and find a suitable pupation site in a crack
or crevice or on a flat surface that may be several meters from the water. Here, they spin a double-walled
cocoon in which they pupate, the outer wall having a
distinctive meshlike construction. After several days,
pharate adults use pupal mandibles to cut their way
out of the cocoon and emerge. Adults feed on nectar
and apparently live no more than a few weeks. Most
species overwinter as larvae on sponges, but some may
overwinter as prepupae in the pupal cocoon.
Larvae have no special adaptations for the aquatic
environment. They obtain oxygen from the water
through their cuticle and swim by repeatedly arching
their abdomen forward while in a vertical position and
snapping it back. Larvae are rarely collected by standard sampling techniques. Substantial numbers can be
collected only by collecting sponges and removing larvae that crawl from them as they dry. Rearing has been
accomplished, but a suitable colony of freshwater
sponges must be maintained and a terrestrial habitat
for pupation must be provided.
C. Aquatic Lepidoptera — Aquatic Caterpillars
Larvae in several genera of this large, primarily terrestrial order, are associated with aquatic habitats.
Some feed on emergent parts of aquatic macrophytes
and others mine stems of these plants. Still others feed
on submerged parts of plants attached to other substrates, and only these will be considered here. The 17
aquatic genera in which aquatic larvae occur are all in
686
W. L. Hilsenhoff
the family Pyralidae, and almost all are in the subfamily Nymphulinae.
Aquatic caterpillars have three pairs of thoracic legs
and pairs of short prolegs ringed with hooklike crochets
on abdominal segments 3 – 7 (Fig. 103). They are not
likely to be confused with any other aquatic insect larvae, but often it is not possible to distinguish them from
Lepidoptera that are inadvertently collected from emergent vegetation. Only those that have numerous filamentous gills covering their body and those that live in
portable cases made from the aquatic vegetation that
they inhabit, can be recognized with certainty as being
aquatic. Larvae that live in cases are so well camouflaged that they often escape notice, even after capture
with a net. Some species may become sufficiently abundant to have an impact on aquatic plant communities.
While adults have been well studied and larvae of many
of the species have been reared and described, reliable
keys to species of larvae have not been constructed because larvae of many species remain unknown.
Larvae inhabit a wide variety of permanent aquatic
habitats. Some inhabit rocks in rapid water of larger
streams where they feed on diatoms and other algae.
Others inhabit aquatic macrophytes or duckweed
(Lemna) in ponds or along margins of streams and
lakes. Here, they feed on the plants they inhabit and
make cases from them. Respiration is cutaneous, and is
enhanced by numerous filamentous gills in larvae that
inhabit rocks in streams and also in some lentic casebuilders. Larvae of at least one species have hydrofuge
hairs enabling them to maintain a plastron that is used
as a physical gill. Pupation of aquatic species is in a cocoon, usually in the larval habitat. Emergence occurs after pupae swim or crawl to the surface of the water; legs
of species that swim have long swimming hairs. Adults
are generally nondescript small moths that hold their
wings rooflike over their body and remain in the vicinity of their larval habitat. Adults of stream species crawl
under the water to oviposit on rocks, while those of
most lentic species deposit rows of eggs just below the
water surface on the preferred food plant. Most species
are univoltine or bivoltine and have five larval instars.
D. Aquatic Coleoptera — Water Beetles
Only about 3% of Coleoptera species have an
aquatic stage, but because this is the largest insect order they represent a significant segment of the aquatic
insect fauna. About 1450 aquatic species in 19 families
have been found in North America. There are also several species of riparian Coleoptera that occasionally
may be collected while sampling margins of aquatic
habitats. In eight families both larvae and adults are
aquatic in almost all species, and in nine others either
all larvae or all adults are aquatic. A few families with
mostly terrestrial species also have some species with
larvae and/or adults that are adapted for living in the
aquatic environment.
Three suborders of Coleoptera have aquatic representatives. Adephaga is represented by five families in
which all species are aquatic both as larvae and adults.
These families are often referred to as the “Hydradephaga”. The suborder Myxophaga is represented in
North America by a single aquatic species, Hydroscapha
natans, in which both larvae and adults are aquatic.
Several families of Polyphaga have some species with
one or more aquatic stages, but only in a few families do
most species have a tie to the aquatic environment.
Most aquatic beetles can be collected with an
aquatic net by using standard collecting techniques.
However, larger Dytiscidae and Hydrophilidae are ineffectively sampled with nets, and bottle traps must be
relied on to capture substantial numbers of these larger
beetles and also species that are nocturnal (Hilsenhoff,
1987, 1991). Adult Gyrinidae, which mostly swim on
the surface in deeper water, are seldom captured without special effort. In late summer, large schools often
appear in shallow bays of lakes and streams, and by
herding them toward shore and sweeping through the
school, large numbers often can be captured. In the
north, where ponds and lakes freeze, lentic as well as
lotic gyrinids congregate in streams in the fall to overwinter. Here, very large numbers of mixed species can
be collected with a net from undercut banks where
there is a current and the water is at least 0.5 m deep.
Some overwintering lentic species of Dytiscidae, Haliplidae, and Hydrophilidae also may be found along
stream banks in autumn and early spring. Large numbers of Hydrophilidae, Helphoridae, and Hydrochidae
adults can often be collected by black-light traps on
warm nights in spring and summer.
Adult water beetles range from 1 – 40 mm in length
and vary greatly in structure. All are characterized by
chewing or biting mouthparts and shell-like mesothoracic wings that are called elytra (Fig. 108). They have
functional spiracles and rely on atmospheric oxygen for
respiration. Adult Adephaga carry a bubble of air under their elytra, which is renewed at the surface by
swimming or crawling (Amphizoidae) to the surface
and breaking the surface film with the tip of the abdomen. This air supply must be frequently renewed,
except when water temperatures are cold and metabolism is slowed, or when the water is well oxygenated.
Hydroscaphidae adults (Myxophaga) crawl to the surface to renew their air supply. Elmidae and Dryopidae
adults (Polyphaga) usually live in oxygen-rich water
and maintain a plastron of air that is carried on hydrofuge hairs. The plastron acts as a physical gill, with
687
17. Diversity and Classification of Insects and Collembola
oxygen for respiration diffusing into it from the water,
and carbon dioxide from respiration diffusing out into
the water. Other Polyphaga adults (Helophoridae, Hydrochidae, Hydrophilidae, Hydraenidae, and Curculionidae) also maintain an air supply on hydrofuge
hairs, but must renew the air by swimming (most Hydrophilidae) or crawling (Curculionidae, Hydraenidae,
Helophoridae, Hydrochidae, and some Hydrophilidae)
to the surface when the water contains too little oxygen
for plastron respiration. Unlike Adephaga, adult
Polyphaga break the surface film with their antennae.
Larvae of aquatic Coleoptera vary greatly in size
and general morphology. All have three pairs of thoracic legs, chewing or biting mouthparts, and are relatively well sclerotized, especially in the later instars.
Some have lateral filaments and resemble larvae of
Megaloptera. Most larvae of aquatic Coleoptera rely on
cutaneous respiration, which may be enhanced by gills,
although some larger Dytiscidae and Hydrophilidae
larvae obtain oxygen at the water surface through functional posterior spiracles. Larvae of Chrysomellidae and
Noteridae are able to utilize oxygen from plant tissues,
and by this mechanism are able to pupate under water.
It is also likely that larvae and adults of Celina (Dytiscidae) obtain oxygen from plants (Spangler 1973;
Hilsenhoff, 1994). Most Coleoptera larvae lack spiracles in the early instars, but are equipped with a full
complement of spiracles in the final instar, which allows
them to leave the water and pupate in cells on land.
Adephaga adults (except Amphizoidae) and many
Hydrophilidae adults are adapted for swimming by
having long setae or swimming hairs on at least their
metathoracic legs or by having greatly flattened legs
(Gyrinidae). Some larvae of Dytiscidae are also adapted
for swimming by having long setae on their legs, but
larvae of most aquatic Coleoptera crawl on vegetation
or other substrates. Gyrinidae larvae, which mostly
crawl on vegetation, can swim rapidly with an undulating motion when disturbed.
Much of the taxonomy of North American water
beetles was completed more than 50 years ago, but
some genera need revision and several species remain
undiscovered. Adults of almost all species can be reliably identified, but their distribution remains poorly
defined in North America because in many areas
aquatic beetles have not been adequately studied.
Unlike adults, very few larvae can be identified to
species and a few cannot even be identified to genus. In
most genera, larvae of too few species are known to
permit development of reliable keys. Fortunately, the
European fauna is much better known, and the European literature gives us insight into probable life cycles,
larval and adult habitats, overwintering sites, feeding
habits, and larval taxonomy. Development of keys and
descriptions for aquatic Coleoptera larvae remains a
pressing need. To accomplish this, larva – adult associations must be obtained through laboratory rearing. The
most promising method for doing this is to capture
mated females, or male – female pairs, allow females to
oviposit, and then rear larvae that hatch from the eggs.
By using this ex-ovo method (Alarie et al., 1989), all
larval instars can be associated with the adult.
Because of our inability to identify larvae, life histories of most North American species remain poorly
known. This includes our knowledge of adult habits,
especially such things as feeding, dispersal, overwintering, and estivation. From the occurrence of teneral
adults and collection records for adults and larvae (if
identified), life cycles and overwintering sites have been
postulated for many species of Wisconsin Dytiscidae,
Helophoridae, Hydrochidae, and Hydrophilidae
(Hilsenhoff (1992, 1993 a-c, 1994, 1995a,b,d). Much
research is needed before we will understand the bionomics of the many species of aquatic Coleoptera in
North America.
References to species keys for the identification of
Coleoptera adults and larvae are provided in Table VII.
Keys to families of freshwater Coleoptera adults and
TABLE VII Literature References to Species Keys for
Identification of Coleoptera Adults and Larvae
Regional keys to adults that include more than one family
California – Leech and Chandler (1956)
Florida – Young (1954); Epler (1996)
Maine – (Adephaga and Hydrophilidae) Malcolm (1971)
North Dakota – Gordon and Post (1965)
North and South Carolina — Brigham (1982c)
Pacific Northwest — Hatch (1953, 1965)
Keys to adults listed by family or superfamily
Amphizoidae — Kavanaugh (1986)
Dryopoidea — Brown (1972); (Louisiana) Barr and Chapin (1988);
(Wisconsin) Hilsenhoff and Schmude (1992)
Dytiscidae — (Alberta) Larson (1975); (Canada and Alaska)
Larson et al. (2001); (Virginia) Michael and Matta (1977);
(Utah) Anderson (1962); (Wisconsin) Hilsenhoff (1992,
1993a, b, c, 1994, 1995a)
Acilius — Hilsenhoff (1975)
Agabus — Fall (1922); Larson (1989, 1991, 1994, 1996, 1997);
Larson and Wolfe (1998); (Pacific Coast) Leech (1938)
Celina — Young (1979a)
Colymbetes — Zimmerman (1981)
Copelatus — Young (1963a)
Coptotomus — Hilsenhoff (1980)
Desmopachria — Young (1981a, 1981b)
Dytiscus — Roughley (1990)
Graphoderus — Wallis (1939)
Heterosternuta — Matta and Wolfe (1981)
Hydaticus — Roughley and Pengelly (1981)
Hydroporus — Gordon (1969, 1981)
(Continues)
688
TABLE VII
W. L. Hilsenhoff
(Continued)
Hydrovatus — Bistrom (1996), Young (1956, 1963b)
Hygrotus — Anderson (1970, 1975, 1983)
Ilybius — Larson (1987)
Laccophilus — Zimmerman (1970)
Laccornis — Wolfe and Roughley (1990)
Liodessus — Larson and Roughley (1990); Miller (1998)
Lioporeus — (as Falloporus) Wolfe and Matta (1981)
Matus — Young (1953)
Nebrioporus — Shirt and Angus (1992); (as Deronectes)
Zimmerman and A.H. Smith (1975)
Neoporus — (as Hydroporus) Fall (1923); Wolfe (1984)
Oreodytes — Zimmerman (1985); Alarie (1993)
Rhantus — Zimmerman and R. L. Smith (1975)
Sanfilippodytes — (as Hydroporus) Fall (1923)
Stictotarsus — (as Deronectes) Zimmerman and
A. H. Smith (1975)
Thermonectus — McWilliams (1969)
Elmidae
Dubiraphia — Hilsenhoff (1973)
Optioservus — White (1978)
Stenelmis — Schmude (1992), (1999)
Gyrinidae — (Minnesota) Ferkinhoff and Gundersen (1983);
(Quebec) Morrissette (1979); (Wisconsin) Hilsenhoff (1990)
Dineutus — Hatch (1930); Wood (1962)
Gyrinus — Oygur and Wolfe (1991)
Haliplidae — (Minnesota) Gundersen and Otremba (1988);
(Virginia) Matta (1976); (Wisconsin) Hilsenhoff and
Brigham (1978)
Haliplus — Wallis (1933)
Peltodytes — Roberts (1913)
Helophoridae — Smetana (1985); (Pacific Northwest)
McCorkle (1965)
Hydraenidae — Perkins (1980)
Hydrochidae — Hellman (1975)
Hydrophilidae (includes Helophoridae and Hydrochidae) —
(Canada and Alaska) Smetana (1988); (Illinois) Wooldridge
(1967); (Michigan) Willson (1967); (Mississippi) Testa and
Lago (1994); (Nevada) LaRivers (1954); (Pacific Northwest)
Miller (1965); (Virginia) Matta (1974); (Wisconsin) Hilsenhoff
(1995b, d)
Anacaena — (Europe) Berge Henegouwen (1986)
Berosus — Van Tassell (1966)
Crenitis — Winters (1926)
Cymbiodyta — Smetana (1974)
Enochrus — Gundersen (1978)
Hydrochara — Smetana (1980)
Laccobius — Cheary (1971); Gentili (1985, 1986)
Paracymus — Wooldridge (1966)
Tropisternus — Spangler (1960)
Noteridae
Hydrocanthus — Young (1985)
Suphisellus — Young (1979b)
Keys to larvae listed by family
Chrysomelidae — (Michigan) Hoffman (1939)
Dytiscidae — (Wisconsin) Hilsenhoff (1992, 1993a, b)
Heterosternuta — Alarie (1992)
Hydroporus — Alarie (1991)
Hygrotus — Alarie et al. (1990)
Oreodytes — Alarie (1997)
Ptilodactylidae
Anchytarsus — Stribling (1986)
larvae appear in Sections V.H.1 and V.H.2. Information on suborders and the biology, habitat, and identification of adults and larvae in each family is presented
in the following.
1. Families of Aquatic Coleoptera — Suborder
Adephaga
In the five aquatic families of this suborder (Hydradephaga), both larvae and adults of all species are
aquatic. Eggs are laid singly or in masses, mostly on
objects beneath the water surface. Larvae have three instars, and except for at least some Noteridae, pupation
takes place in terrestrial cells near the larval development site. Almost all first and second instar larvae lack
spiracles and rely on cutaneous respiration for oxygen.
A few have functional posterior spiracles that they use
to obtain oxygen at the water surface. Third instar larvae, which have functional spiracles on abdominal segments and the mesothorax, also rely mostly on cutaneous respiration until they leave the water to pupate.
Life cycles are typically univoltine, although bivoltine
and semivoltine life cycles have been recorded for some
species. Some species complete larval development
rapidly, requiring only a few weeks; others require several weeks, and those with semivoltine life cycles may
need almost two years to complete larval development.
Adults are long-lived, with records of unmated beetles
living for several years, but typically most adults live
several months and die after their final mating and
oviposition, which is usually in spring.
a. Amphizoidae (1 Genus, 3 Species) “Trout-stream
beetles” are restricted to mountain streams in northwestern North America. Here, both larvae and adults
can be found near the water edge, crawling on plant
roots, wood, or debris at or just beneath the water surface. Larvae are characterized by short, broad urogomphi and flattened lateral projections on each segment
(Fig. 120). Adults are relatively large (12 – 14 mm long)
and broad, with a narrow thorax and no swimming
hairs on their legs (Fig. 108). Larvae cannot swim and
adults swim very poorly; both usually remain in contact
with the substrate. Third instar larvae frequently stay in
the splash zone and enter the water only to capture
prey. Both larvae and adults feed almost exclusively on
Plecoptera larvae. Because their habitat is well-oxygenated, adults may remain submerged for prolonged
periods, their air bubble acting as a physical gill. The
life cycle is 1 year, with eggs laid on debris in the splash
zone in late summer and pupation occurring the following summer, probably in cells in the stream bank.
b. Dytiscidae (48 Genera, 600 Species) The
“predaceous diving beetles” are the largest family of
17. Diversity and Classification of Insects and Collembola
water beetles, with more than 550 species distributed
throughout North America. Adults range from 2 to
40 mm in length and are characterized by their ovate
appearance, rounded sternum and dorsum, and by a
usually spear-shaped prosternal process that projects
back to the mesocoxae (Fig. 106). The beetles swim by
moving both metathoracic legs backward simultaneously. Larvae are generally elongate, and range from 3
to 60 mm in length at maturity. Most have paired terminal appendages (urogomphi) of various lengths (Fig.
122). They have relatively elongate legs, which in several genera are modified by long hairs for swimming.
Adults and larvae of most species are lentic, inhabiting all types of permanent and temporary habitats,
and showing a preference for the shallow, vegetated
margins of ponds, marshes, bogs, and swamps. They
are most abundant in habitats that do not contain significant numbers of insectivorous fish. Most species
breed in well-defined habitats, but adults may fly to
other habitats to feed or overwinter. Adults and larvae
of lotic species are found mostly under stream banks
and tend to remain in the habitat in which they breed.
Typically adults overwinter, mate in the spring,
oviposit, and die. Eggs are laid in or on vegetation beneath the water surface, or on other objects below or
just above the surface. The larvae develop over a period of a few to several weeks, then leave the aquatic
habitat to pupate in cells they construct in the soil of
protected areas nearby. The pupal stage lasts 5 – 14
days, after which the adult emerges and usually re-enters the aquatic habitat. Most species are univoltine,
but bivoltine life cycles occur, especially in the south,
and there is evidence northern species that breed in
cold-water habitats are semivoltine. However, life histories of many species remain poorly known, and there
are variations in the life cycle described above. There is
ample evidence for estivation of adults during dry periods in the summer and also evidence that several northern species overwinter as diapausing eggs. Many lotic
species and a few lentic species overwinter as larvae,
even in northern areas that freeze. While most overwintering adults are found in ponds, lakes, or streams,
some pass the winter in terrestrial habitats and enter
their breeding sites late in the spring. Both adults and
larvae are predators, feeding primarily on other invertebrates and small vertebrates. Adults of most species
are also scavengers.
c. Gyrinidae (4 Genera, 60 Species) Adult “whirligig
beetles” are small- to medium-sized beetles (3.5 – 14.0
mm long). They are widespread and often abundant.
Most frequently, they are seen resting or swimming in
groups of a few to several hundred on the water surface
in areas protected from the wind, especially in late
689
summer. However, they can, and often do, swim under
the surface. Their greatly flattened legs and two pairs of
eyes are distinctive. Larvae are also distinctive, having
fringed projections on each abdominal segment posteriorly (Fig. 119). They swim with a vertical undulating
motion when disturbed.
While adults and larvae of most species are lentic,
those of several species are lotic, and adults of most
lentic species frequent larger streams in autumn, or in
late summer when it is dry. In streams, adult gyrinids
will often escape the current by crawling onto vegetation just above the water. In northern regions, where
ponds and lakes freeze, adults of most lentic species fly
to larger streams or lakes to overwinter. They return to
breeding sites in early spring, as soon as air temperatures have warmed enough to permit flight. All species
tend to inhabit deeper water than other Hydradephaga,
with larvae being found mostly among submerged vegetation in large ponds or rivers, or in littoral zones of
lakes and impoundments. Larvae are predators, feeding
mostly on other invertebrates, while adults are scavengers on dead animals or predators of small invertebrates in the surface film. Most species are probably
univoltine, with adults overwintering and larvae developing during the summer. Eggs are laid mostly on submerged aquatic vegetation. Pupation is in cocoons on
emergent vegetation, or on terrestrial vegetation adjacent to the larval habitat.
d. Haliplidae (4 Genera, 70 Species) “Crawling
water beetles” are small (2.5 – 5.0 mm long) yellowish
to orangish beetles with black blotches and spots, apically narrowed elytra, and a narrowed thorax. Their
large metacoxal plates are distinctive (Fig. 105). They
are often abundant in shallow lentic and lotic vegetation-choked habitats. In spite of their common name,
they have fringed tarsi and tibiae and are able swimmers. The larvae, which crawl about on vegetation, are
either elongate with a long terminal segment (Fig. 123),
or have numerous long dorsal projections (Fig. 124).
Life cycles of most species are probably univoltine,
with eggs being deposited on or in filamentous algae,
or other vegetation. Both larvae and adults are herbivores, feeding on algae or macrophytes. Pupation takes
place in cells in moist soil near the larval development
site. Most species overwinter in the water as adults, but
some adults and larvae are known to overwinter in terrestrial sites adjacent to the water.
e. Noteridae (5 Genera, 18 Species) This small
family, frequently referred to as “burrowing water beetles,” is generally southern in its distribution and rarely
found as far north as Canada or in the west. Adults are
small (1.2 – 5.5 mm long) and resemble Dytiscidae, but
690
W. L. Hilsenhoff
are usually more attenuate apically than most Dytiscidae. They have a flat, V-shaped metasternal plate and a
truncate, instead of a spear-shaped or rounded, prosternal process (Fig. 111). Like Dytiscidae, adults are
mostly predators that live in close association with
aquatic vegetation. The larvae, which are poorly
known, have short legs, short urogomphi, and telescoping body segments (Fig. 121). Their morphology
suggests that they are omnivores, which live among
roots of aquatic vegetation. Larvae of a European
species obtain oxygen from plant tissues and pupate in
a cocoon on the plant roots, the pupae also obtaining
oxygen from the plant. Life cycles are probably one
year, with adults overwintering in the water.
2. Families of Aquatic Coleoptera — Suborder
Myxophaga
a. Hydroscaphidae (1 Genus, 1 Species) The only
North American species occurs from the western
United States, north to Idaho. The fusiform shape,
truncate elytra, and very small size (1.5 mm) of adult
“skiff beetles” is distinctive (Fig. 118). Larvae and
adults are found most commonly on algae over which a
thin film of water flows, but adults have also been collected in deeper water. Both adults and larvae are herbivores, feeding on algae. They tolerate a wide range of
water temperatures, and have been found in thermal
springs as warm as 46°C and in cold streams that frequently freeze. Females lay single, well-developed eggs
on algae, and pupation also takes place on algae at the
air – water interface. Larval respiration is aided by distinctive pairs of balloon – like spiracular gills on the
prothorax, and first and eighth abdominal segments.
Adults, which may remain submerged for long periods,
retain an air bubble that acts as a physical gill.
3. Families of Aquatic Coleoptera — Suborder
Polyphaga
There are four families in which both aquatic
adults and larvae are found. In Elmidae, adults and larvae of almost all species are aquatic, while in Hydrophilidae most species have aquatic adults and many
also have aquatic larvae. Curculionidae has several
species with aquatic adults and a few also with aquatic
larvae, while Chrysomelidae has several species with
aquatic larvae and two also with aquatic adults. In
Dryopidae, Hydraenidae, Helophoridae, and Hydrochidae all adults are aquatic, and in Lutrochidae,
Psephenidae, Ptilodactylidae, and Scirtidae almost all
larvae are aquatic. Larvae in at least one species of
Lampyridae are also aquatic. Additional families of
Polyphaga have species that are associated with water,
but not adapted for living in it, and these families will
not be treated here.
Five families (Dryopidae, Elmidae, Lutrochidae,
Psephenidae, and Ptilodactylidae) form a natural group
of beetles in which almost all species have at least one
aquatic stage. They formerly were placed in the superfamily Dryopoidea and will be discussed first (Section
3.a – e), although they now are included in a larger superfamily (Byrrhoidea) (Lawrence and Britton, 1991).
They differ from most other aquatic beetles by generally having more larval instars, longer life cycles, and
longer adult lives in species with aquatic adults.
Aquatic adults and larvae are unable to swim, which
confines them to well-oxygenated habitats. Aquatic larvae obtain oxygen from the water through their integument, usually with the aid of gills. Aquatic adults have
a permanent air bubble (plastron) that is carried on hydrofuge hairs and acts as a physical gill. Terrestrial
adults of species with aquatic larvae typically inhabit
the splash zone on rocks that project from their aquatic
habitats, and they also use a plastron to remain submerged for relatively long periods when ovipositing. In
Dryopoidea that have been studied, five-to eight-larval
instars have been recorded, but there is disagreement
about the actual number of larval instars, with six being suggested for many species. In Adephaga,
Helophoridae, Hydrochidae, Hydrophilidae, and Hydraenidae there are only three larval instars. Larval development probably takes about 2 years for most
species of “Dryopoidea”, with 1 – 3-year development
times having been suggested for various species. The
shorter times are for species in warmer and more
southern streams. Aquatic adults typically live a year or
more, and some may live several years; terrestrial
adults of species with aquatic larvae typically live only
a few days to a week. To date, studies of life histories
have been mostly on Elmidae and Psephenidae; additional studies are needed. Brown (1987) reviewed the
biology of these riffle beetles.
Larvae of “Dryopoidea” are poorly known, and
identification beyond genus is usually not possible.
Brown (1972) summarized what was known about the
distribution and taxonomy of Dryopoidea in the United
States, and provided species keys for adults and generic
keys for larvae. Since his publication, the aquatic
Limnichidae have been recognized as a separate family,
Lutrochidae. Several new species have been discovered
recently, and revisions are needed for some genera.
a. Dryopidae (5 Genera, 14 Species) In Helichus,
the most common and widespread genus of “long-toed
water beetles,” only adults are aquatic, living among
debris and on decaying wood in streams. Adults in
other genera are also probably aquatic, with those in
two genera being lotic and those in the other two genera lentic. Larvae and adults resemble those of other
17. Diversity and Classification of Insects and Collembola
Dryopoidea. Adults are larger than the more abundant
Elmidae and have a pectinate antenna (Fig. 116). Larvae are elongate and lack gills in the operculate caudal
chamber. Little is known about life histories of species
in this family.
b. Elmidae 27 Genera, 101 Species) Commonly referred to as “riffle beetles,” Elmidae are widespread
and often abundant. Both larvae and adults are usually
aquatic and often occur together; in a few species,
adults are riparian. Adults are less than 4.5 mm long,
smaller than Dryopidae adults, and have filiform or
slightly clubbed antennae (Fig. 117). The sclerotized
larvae are elongate, rounded in cross section, and have
a ventral caudal operculum that closes a chamber containing hooks and numerous filamentous gills (Fig.
132). Both adults and larvae are found mostly in
streams, where they inhabit a variety of substrates, including gravel riffles, algae laden rocks, aquatic macrophytes, and decaying wood. Adults and larvae of some
species may also occur on these same substrates in
spring ponds or along wave-swept shores of lakes. Larvae and adults are herbivores – detritivores, feeding on
algae, decaying wood, and detritus. When larvae complete their development they leave the water and pupate in cells in protected areas on the adjacent shore.
Upon emergence, adults disperse widely and frequently
are captured in light-traps. Once they find a suitable
aquatic habitat, they rarely if ever fly again, but stream
species may move downstream by drifting in the current. Most adults probably live a year or more, and a
semivoltine life cycle seems probable for most species
in the north.
c. Lutrochidae (1 Genus, 3 Species) Larvae of
Lutrochidae occur on calcareous encrustations in hardwater streams in eastern, central, and western United
States. They closely resemble larvae of Elmidae, but the
last abdominal tergum is rounded apically (Fig. 134)
instead of being emarginate (Fig. 133), and thoracic
sterna are membranous. They apparently feed on periphyton, but little is known about the bionomics of this
family. Adults are riparian, inhabiting splash zones of
larger rocks that project from the aquatic habitat.
d. Psephenidae (5 Genera, 14 Species) “Water pennies” are flat, rounded larvae (Fig. 128) that occur on
rocks and wood in streams from Texas and Arizona
north to British Columbia, and east of the Great Plains;
they may also be found on wave-swept shores of lakes.
In two genera (Psephenus and Eubrianax) larvae lack
the ventral caudal operculum of other aquatic dryopoid
larvae, but have ventral abdominal gills that aid in respiration. The remaining genera are in the subfamily Eu-
691
briinae, which is sometimes considered a separate family. Their larvae lack exposed ventral gills, possess a
ventral caudal operculum, and have the lateral margins
of each segment distinctly separated (Fig. 127). Pupation of most species occurs above the water line on
rocks near the larval habitat, but at least one species is
known to pupate under water. Adults are riparian, typically being found in splash areas of projecting rocks.
Most species probably have life cycles of 1 – 2 years.
e. Ptilodactylidae (3 Genera, 3 Species) Larvae
have been collected from streams and springs in
California and Nevada, and from Georgia to New
York, but not in central North America. They are elongate, lack a ventral caudal operculum, and have gills either on the first seven abdominal sterna or on the ninth
sternum (Fig. 131). Very little is known about their life
histories. Adults of aquatic species are riparian.
f. Chrysomellidae (6 Genera, about 70 Species) In
this very large terrestrial family there are several species
in which larvae and adults feed on emergent aquatic
vegetation, but there is only one subfamily (Donaciinae) in which aquatic larvae feed on submerged portions of aquatic plants. They are adapted for underwater existence by having a pair of caudal spiracular
spines that they insert into plant tissues to obtain oxygen, and they can be recognized by these conspicuous
spines (Fig. 130). Larvae pupate on underwater portions of plants, with pupae also obtaining oxygen from
plants for respiration. Adults of at least two species are
also aquatic, feeding on plant stems.
g. Curculionidae (20 Genera, about 150 Species) In
this very large terrestrial family, adults of species in
several genera feed on aquatic plants, both above and
below the water surface, and there is no easy way to
distinguish truly aquatic species. Adult Curculionidae
are readily distinguished by their unique head, which is
elongated into a snout, and by their geniculate antennae (Fig. 104). Larvae are grublike, without distinct
legs; most mine leaves and other plant tissues and are
not aquatic. Aquatic adults crawl on underwater portions of plants and carry with them a plastron for respiration; a few species are capable of swimming. Most
aquatic species are nocturnal, and difficult to find during the day, when they apparently hide in the bottom
substrate. Life histories are poorly known.
h. Hydraenidae (3 Genera. 82 Species) Adult
“minute moss beetles” are aquatic, living mostly in extremely shallow water along margins of streams, but
also occurring along margins of lentic habitats. Adults
resemble Hydrophilidae, and once were considered
692
W. L. Hilsenhoff
part of that family. They are much smaller (2 mm
long) than most Hydrophilidae and have five segments
in the antennal club instead of three. Larvae are riparian, developing and pupating in moist soil adjacent to
the water. Adults cannot swim, and float upside down
in the surface film when dislodged from the substrate
on which they live. Because of their very small size they
are easily overlooked. Their life history and feeding
habits are poorly known.
i. Helophoridae (1 Genus, 41 Species) Helophorus
was previously included in the family Hydrophilidae,
but a study by Archangelsky (1998) convinced me to
include it in a separate family. Adults are aquatic, but
are poorly adapted for swimming and mostly crawl
about on aquatic vegetation in a variety of lentic habitats or along margins of streams. Larvae inhabit riparian habitats adjacent to aquatic habitats where adults
are found. Adults are small, elongate beetles that are
readily recognized by the seven longitudinal grooves
and six raised areas on the pronotum (Fig. 114). The
univoltine life cycles are variable, with adults or eggs
overwintering in riparian habitats, depending on the
species (Hilsenhoff, 1995b).
j. Hydrochidae (1 Genus, 19 Species) Hydrochus
also was included in Hydrophilidae, but is now considered to be in a separate family (Archangelsky, 1998).
Like Helophoridae, adults are aquatic, but swim
poorly and mostly crawl about on the aquatic vegetation of shallow lentic habitats. Occasionally they occur
along margins of streams. Life cycles are univoltine,
with adults overwintering in terrestrial habitats, flying
back to aquatic habitats in spring, and ovipositing in
adjacent riparian habitats where the larvae develop.
Adults are readily identified by their small size and
a very narrow pronotum that is covered with small
granules and / or punctures (Fig. 115).
k. Hydrophilidae (19 Genera, 160 Species) Adults
and larvae of this large and abundant family mostly inhabit shallow lentic habitats, although a few species are
found in deep water and others are strictly lotic. In several genera only adults are aquatic, and in one subfamily (Sphaeridiinae) none of the life stages are aquatic.
While commonly called “water scavenger beetles,” all
aquatic larvae are predators and adults feed on algae
and perhaps small animals, as well as scavenging.
Adults range in size from 1.5 to 40 mm and somewhat
resemble Dytiscidae, but are flat ventrally, have clubshaped antennae (Fig. 113), and very long maxillary
palps. The predatory larvae have large, conspicuous
mandibles and relatively long antennae and maxillary
palps (Fig. 125).
Adults in many genera mostly crawl on aquatic
vegetation and are poorly adapted for swimming. In
genera with adults that swim, beetles have fringes of
long setae on their meso- and metathoracic legs, which
they move alternately instead of in unison as in the
Dytiscidae. While adults of most species can be readily
collected with an aquatic net, those of larger species
that swim rapidly are most effectively collected with
bottle traps. Most species are univoltine, with adults
being the normal overwintering stage. Adults of several
species overwinter in terrestrial habitats, while others
that inhabit shallow lentic habitats may fly to streams
or larger lentic habitats in late summer and autumn to
overwinter.
l. Lampyridae Larvae of at least one species of
“firefly” have been collected from shallow lentic and
semilotic habitats. They are readily recognized by their
flat, platelike thoracic nota that mostly or completely
conceal the head from above, and by gills ventrally on
the last abdominal segment (Fig. 129). Larvae are
predaceous, feeding on snails or other insects.
m. Scirtidae (8 Genera, 35 Species) Although
adults are terrestrial, larvae of “marsh beetles” are
aquatic and often common in marshes, swamps, margins of shallow ponds, tree holes, and a variety of other
lentic habitats where they crawl about on the aquatic
vegetation. Their elongate filiform antennae are unique
among aquatic Coleoptera larvae (Fig. 126). They are
apparently detritivores, with little being known about
their life history.
E. Aquatic Diptera — Flies and Midges
Diptera is a large, mostly terrestrial order, but larvae and pupae of numerous species are aquatic, and it
is the dominant order of insects in the aquatic environment. Diptera larvae inhabit all types of aquatic environments, and because of very short development periods in many species, they are able to utilize ephemeral
aquatic habitats as well as permanent ones. About
40% of all species of aquatic insects belong to this order, and at least a third of the aquatic Diptera species
are in one family, Chironomidae. It is impossible to accurately estimate the number of species of aquatic
Diptera, because in most families only terrestrial adults
can be identified and in many families it is not possible
to know which species have aquatic larvae. It is also
difficult to estimate the relatively large number of
aquatic species that probably remain undiscovered and
undescribed.
The order is divided into two suborders, Nematocera and Brachycera (Agriculture Canada, 1981), with
17. Diversity and Classification of Insects and Collembola
Nematocera dominating the aquatic fauna. Formerly a
third suborder, Cyclorrhapha, was recognized; it is
now included in Brachycera as the infraorder Muscomorpha. In Nematocera there are several families in
which all or almost all species have aquatic larvae and
pupae. In Brachycera most families that have species
with aquatic larvae also have large numbers of terrestrial or riparian species. Larvae of the infraorder Muscomorpha are often referred to as cyclorrhaphous
Brachycera, with the remaining Brachycera being called
orthorrhaphous Brachycera.
Diptera larvae lack segmented thoracic legs, and
thus can be readily distinguished from larvae of other
aquatic insects. In Nematocera larvae, a completely
sclerotized, relatively round head capsule is present (except in most Tipulidae), and the mandibles, which usually have subapical teeth, move laterally. In Brachycera
larvae the head is reduced to an internal skeleton (cyclorrhaphous Brachycera) or poorly formed and not
rounded (orthorrhaphous Brachycera), and the
mandibles (mouth hooks) move vertically and lack subapical teeth.
Diptera larvae have many adaptations that allow
them to live in a wide variety of environments. Those
that live in oxygen-rich environments usually rely on
cutaneous respiration, which may be aided by gills that
increase the surface area. In Chironomidae, larvae of
some species have a hemoglobinlike pigment that aids
in oxygen storage. Larvae in many families obtain oxygen at the water surface, aided by short or elongate
caudal respiratory siphons that reach to the surface and
may be retractile. Other larvae have fringes of hairs
surrounding caudal spiracles that allow them to float in
the surface film with their caudal spiracles exposed to
the air. Some larvae repeatedly swim to the surface to
obtain oxygen (Culicidae). Other larval adaptations include papillae for regulation of salt concentration,
sucker disks to anchor lotic forms in rapid currents,
and prolegs, pseudopods, or creeping welts on various
segments to aid in locomotion.
Pupae of Nematocera often have anterior respiratory horns that enable them to obtain oxygen at the
surface; others rely on cutaneous respiration. A few
swim actively to the surface (Culicidae, Chaoboridae)
or pupate just above the water line (Dixidae). Most orthorrhaphous Brachycera pupate on land. Cyclorrhaphous Brachycera and Stratiomyidae pupate in a puparium, which may be terrestrial or float at the water
surface where oxygen is plentiful.
Aquatic Diptera larvae can be readily captured
with an aquatic net as described earlier, and when
placed in a pan with some water will usually swim or
crawl free of the debris, allowing them to be collected.
Active Nematocera larvae are difficult to capture with
693
forceps, and the use of a small loop (30-mm diameter)
of nylon mesh is often advantageous. Such a loop can
be made by bending the end of a piece of thin wire into
a loop at a right angle to the remainder of the wire (the
handle) and gluing a piece of nylon stocking to the
loop. Diptera larvae are relatively easy to rear because
most have short life cycles and pupate in the water.
Lentic species, which require little or no aeration, are
especially easy to rear.
Diptera larvae are not only a very important part
of almost all aquatic ecosystems, but adults in many
aquatic families are important because they transmit
disease or create severe nuisance problems. Culicidae
(mosquitoes) are the most important group from a
public health standpoint because adults transmit diseases of humans, birds, and other animals. Adults of
this family, along with those of Tabanidae (horseflies
and deerflies), Simuliidae (blackflies), and Ceratopogonidae (biting midges) may create severe nuisance problems because they bite humans and other animals.
Nonbiting midges in the families Chironomidae and
Chaoboridae may emerge in such tremendous numbers
from larger lakes that they damage property and create
public health problems by causing severe allergic reactions in some people.
Much research is needed on aquatic Diptera in
North America. Many species remain to be described,
and larvae of most described species remain unknown.
Only in families that are of public health importance
(Culicidae, Tabanidae, Simuliidae) or useful as a biological control (Sciomyzidae) are larvae well enough
known that reliable keys to most species have been developed. Even in these families there are taxonomic
problems, especially with separation of sibling species.
Great strides in larvae taxonomy have been made in recent years, especially in the largest family, Chironomidae, but until much more is accomplished it will not be
possible for us to fully understand the role of Diptera
larvae in aquatic ecosystems. The two-volume Manual
of Nearctic Diptera (1981, 1987) by Agriculture
Canada provides generic keys as well as information
about morphology, biology, behavior, classification,
and distribution.
References to species keys for the identification of
larvae are provided in Table VIII and keys to families
of freshwater Diptera larvae appear in Sections V.I. Information on suborders, and the biology, habitat,
adaptations, and identification of larvae in each family
is presented next.
1. Families of Aquatic Diptera – Suborder
Nematocera
More than half of the families in this suborder
have aquatic representatives, and in 12 families all or
694
W. L. Hilsenhoff
TABLE VIII Literature References to Species Keys
for Identification of Diptera Larvae
Regional keys to larvae that include more than one family
North and South Carolina — Webb and Brigham (1982)
Keys to larvae listed by family
Blephariceridae
Blepharicera — Hogue and Georgian (1986)
Ceratopogonidae — Glukhova (1977)
Chaoboridae — Cook (1956)
Chaoborus — Saether (1970)
Chaoborus — (Schadonophasma) Borkent (1979)
Chironomidae — references since Simpson (1982) and
Wiederholm (1983); (Florida) Epler (1995)
Brillia — Oliver and Rousell (1983)
Cricotopus — Simpson et al. (1983)
Dicrotendipes — Epler (1987)
Endochironomus, Endotribelos, Synendotendipes,
Tribelos — Grodhaus (1987)
Eukiefferiella, Tvetenia — Bode (1983)
Guttipelopia — Bilyj (1988)
Pagastia — Oliver and Roussell (1982)
Paraphaenocladius — Saether and Wang (1995)
Polypedilum — Boesel (1985); Maschwitz and Cook (1999)
Stenochironomus — Borkent (1984)
Tanypodinae — (eastern United States) Roback 1985, 1986a,
1986b, 1987)
Corethrellidae — Cook (1956)
Culicidae — Carpenter and LaCasse (1955); Darsie and Ward
(1981); (Canada) Wood et al. (1979); several regional
keys published prior to 1981; (Florida) Breeland and
Loyless (1982)
Deuterophlebiidae — Courtney (1990)
Nymphomyiidae — Courtney (1994)
Sciomyzidae
Dictya — Valley and Berg (1977)
Elgiva — Knutson and Berg (1964)
Hoplodictya — Neff and Berg (1962)
Pherbella — Bratt et al. (1969)
Sciomyza — Foote (1959)
Sepedon — Neff and Berg (1966)
Tetanocera — Foote (1961)
Simuliidae — (Alabama) Stone and Snoddy (1969); (Alberta)
Currie (1986); (Colorado) Peterson and Kondratieff (1995);
(Michigan) Merritt et al. (1978); (New York) Stone and
Jamnback (1955); (Ontario) Wood et al. (1963);
(Pennsylvania) Adler and Kim (1986); (southeastern
United States) Snoddy and Noblet (1976)
Ectemnia — Moulton and Adler (1997)
Gymnopais, Twinnia — Wood (1978)
Prosimulium — (Canada and Alaska) Peterson (1970)
Simulium — Moulton and Adler (1995); (southern California)
Hall (1974)
Tabanidae — (Arizona) Burger (1977); (eastern North America)
Goodwin (1987); Teskey (1969); Teskey and Burger (1976);
(Illinois) Pechuman et al. (1983); (Louisiana) Tidwell (1973)
Atylotus — Teskey (1983)
Tanyderidae — (Alberta) Exner and Craig (1976)
Thaumaleidae
Thaumalea — (Idaho and California) Gillespie et al. (1994)
almost all species have aquatic larvae and usually also
aquatic pupae. Unless noted otherwise, larvae have
four instars, respiration is through the integument, and
pupation is in the larval habitat.
a. Blephariceridae (5 Genera, 28 Species) The distinctive flattened larvae of “net-winged midges” have
seven apparent body segments and a sucker disk ventrally on the first six segments (Fig. 135). They use
their sucker disks to cling to rocks in very fast water of
western and eastern mountain streams, and also
streams in the northern United States and Canada.
Here, they feed on diatoms and other algae. Larvae
move forward slowly with an undulating motion, or
sideways by alternately moving anterior and posterior
sucker disks. Pupation occurs in cracks and depressions
of rocks near the water surface. Adults live 1 – 2 weeks,
with some females being predators on other midges.
Eggs are laid just above the water on rocks, usually
when water levels are receding. They hatch when
flooded by higher water levels.
b. Ceratopogonidae (20 Aquatic genera, 501 Species)
Aquatic larvae of “biting midges” typically are small,
elongate, and without prolegs (Fig. 151). While many
species are riparian or live in moist terrestrial habitats,
a large number of species occur in aquatic habitats that
include tree holes, marshes, swamps, ponds, all areas of
lakes, and streams. Most larvae are able swimmers and
move about with a serpentine swimming motion. They
are poorly known and most have not been associated
with adults, making it difficult even to know which
species develop in aquatic habitats. Reliable keys to larvae of North American genera have not been developed, although a key to larvae in the U.S.S.R.
(Glukhova, 1977) includes most genera found in North
America. Most larvae are carnivores; others are herbivores or detritivores. Pupae of aquatic species hang in
the surface film by their respiratory horns. Adults of a
few aquatic species bite people and may become annoying pests in some areas; most others feed on small
insects. Kettle (1977) reviewed the biology and ecology
of blood sucking Ceratopogonidae.
c. Chaoboridae (3 Genera, 14 Species) Most larvae
are nearly transparent, except for hydrostatic organs,
causing them to be called “phantom midge larvae.”
They are unique in that their enlarged antennae have
been modified for capturing prey such as insect larvae
and small crustaceans (Fig. 148). They occur in a wide
variety of lentic habitats, and are especially abundant
in the profundal or sublittoral zones of some lakes. In
lakes, third and fourth instar larvae rest in the bottom
mud during daylight hours and prey on zooplankton in
17. Diversity and Classification of Insects and Collembola
the limnetic area at night; earlier instars remain limnetic. They are the only insects frequently found in the
limnetic area of lakes. Larvae also occur in permanent
ponds, spring ponds, vernal ponds, and margins of
swamps. Adults do not feed, but their synchronized
emergences may create severe nuisance problems
around large lakes because adults are highly attracted
to lights. Life cycles are univoltine to multivoltine, depending on species, climate, and habitat. Most larvae
can be identified to species.
d. Chironomidae (208 Genera, 2000 Species) Chironomidae is by far the largest family of aquatic insects, with the number of aquatic species surpassing
that in any other order. The larvae, which are recognized because they usually have anterior and posterior
pairs of prolegs (Fig. 158), are diverse in form and size.
They inhabit all types of permanent and temporary
aquatic habitats, and a few species inhabit semiaquatic
or terrestrial habitats. Larvae are often the dominant
insects in the profundal and sublittoral zones of lakes,
and consequently adults are sometimes called “lake
flies,” although they are most often referred to as
“midges.” In larger lakes, adults of some species may
emerge in such tremendous numbers that they create
nuisance problems along their shores. The short-lived
adults cause allergies in some people, invade factories,
spot the paint on houses, and accumulate in large,
odorous piles.
Larvae are an extremely important part of aquatic
food chains, serving as prey for many other insects and
food for most species of fish. Life cycles are highly variable, with many species being univoltine and many others being bivoltine or multivoltine. In the north, life cycles are longer, with some arctic species requiring
several years to complete development. Feeding habits
also vary widely, with herbivores, detritivores and carnivores all commonly represented. Many larvae, especially predators, are free living, but those of most
species construct loose cases of substrate cemented together with salivary secretions. Most are herbivores
and detritivores that graze on fine particles on the substrate, but some are filter feeders, which construct webs
to filter water that they circulate through their case or
retreat. Larvae of most species are quite tolerant of
lowered levels of dissolved oxygen; some can survive in
areas where oxygen levels are so low that oxygen cannot be detected. Such species are usually red and contain a hemoglobinlike pigment that retains oxygen.
These “blood worms” may become abundant in
sewage lagoons or organically polluted areas of lakes
or streams. The pupal stage is short, with pupae of
most species being quiescent. After completing development, the pupa swims to the surface of the water where
695
emergence occurs. Adults do not feed, and generally
live no more than 2 weeks. The biology of the family
was recently reviewed by Pinder (1986).
In recent years many new species have been named
and great progress has been made in rearing and identifying larvae and pupae, but larvae of too many species
remain unknown to permit reliable species keys to be
constructed for larvae in most genera. A bibliography
of taxonomic literature was compiled by Simpson
(1982), and in 1983 Wiederholm keyed larvae to genus
and species groups, provided comments on identification, ecology, and distribution, and also listed known
species keys. Only keys to species or species groups
that were not included by Simpson or Wiederholm are
listed in Table VIII.
e. Corethrellidae (2 Genera, 5 Species) Until recently, this was a subfamily of Chaoboridae. Larvae occur in small pools, tree holes, or artificial containers in
the southeastern United States north to New Jersey and
west to California. Like Chaoboridae larvae, they have
prehensile antennae, but the antennae are very close together basally and not wide apart as in Chaoboridae.
Little is known about their biology.
f. Culicidae (13 Genera, 172 Species) Because they
bite people and transmit disease, “mosquitoes” have
been studied more thoroughly than any other family of
aquatic Diptera. All species can be identified as adults
or larvae, and life histories and habitats of most species
have been documented. Larvae are readily recognized
by their swollen thoracic area (Fig. 147), the presence of
a caudal respiratory tube (siphon) in most genera, and
their characteristic flip-flop swimming motion. Because
of this swimming motion larvae are commonly called
“wrigglers.” Larvae occur in a variety of shallow lentic
habitats that include tree holes, artificial containers,
catch basins, pitcher plants, swamps, shallow temporary or permanent ponds and marshes, and heavily vegetated margins of lakes and streams. They are not
found in moving water or water subjected to wave action. Larvae obtain oxygen at the water surface through
a caudal siphon, which is very abbreviated in Anopheles
(Fig. 147). Larvae of Coquillettidia and Mansonia attach their spinelike siphon to aquatic plants to obtain
oxygen. Most larvae are active swimmers, and feed on
detritus and microorganisms; larvae in two genera are
predators, mostly on other mosquito larvae. Pupae are
also active swimmers, and obtain oxygen at the surface
through thoracic respiratory horns.
Many species are univoltine, with eggs that diapause over the summer, autumn, and winter, and larval
development and emergence occurring in the spring.
Many others are bivoltine or multivoltine. Although
696
W. L. Hilsenhoff
most species overwinter as eggs, a few overwinter as
larvae and several species overwinter as adults. Most
female adults obtain blood from mammals, birds, reptiles, or amphibians, depending on the species, and may
live several weeks or months. Male mosquitoes do not
feed on vertebrates, instead they obtain nectar from
plants. Culicidae larvae often dominate the insect fauna
of temporary ponds and marshes, especially those that
flood in spring and summer.
g. Deuterophlebiidae (1 Genus, 6 Species) The
unique larvae of “mountain midges” are less than
6 mm long and readily recognized by their elongate,
branched antennae and the presence of seven pairs of
stout ventrolateral prolegs on the abdomen (Fig. 153).
They are found in rapid currents of western mountain
streams. Here, they inhabit the upper surface of lightcolored, smooth rocks that have cracks or depressions,
and are usually at or near the water surface. Pupation
is in the same habitat, usually in depressions in darkercolored rocks. In cold water at higher elevations, there
is one generation per year, while at lower elevations
there may be several generations. Eggs within the pupa
are mature, and when adults emerge, mating and
oviposition occur almost immediately. Adults live only
an hour or two.
h. Dixidae (3 Genera, 42 Species) Larvae of “dixid
midges” are found at or just below the water surface in
lentic habitats or densely vegetated margins of streams.
Here, they characteristically lie bent in a U-shape, often
resting on vegetation. They can be recognized by pairs
of prolegs on the first one or two abdominal segments
(Fig. 154). Larvae obtain oxygen from the air through
caudal spiracles and feed on microorganisms and detritus in the surface film. Pupation occurs just above the
water surface on vegetation. Life histories are poorly
known; most species are probably multivoltine.
i. Nymphomyiidae (1 Genus, 2 Species) The small,
slender larvae are recognized by pairs of ventrally projecting, slender, two segmented prolegs on abdominal
segments 1 – 7 and 9 (Fig. 152). They are rare, and in
North America have been found only in Quebec, New
Brunswick, and Maine. Larvae occur mostly on mosscovered rocks in small, rapid, cold, spring-fed streams,
where they apparently feed on diatoms and other algae.
Species are probably bivoltine, with emergences in
spring and late summer.
j. Psychodidae (6 Aquatic genera, 67 Species) Most
species of “moth flies” develop in semiaquatic or moist
terrestrial habitats, but larvae of several species occur
in aquatic habitats. Aquatic larvae can be recognized
by their small size and secondary annulations on thoracic and abdominal segments, which often also have
sclerotized dorsal plates on many annuli (Fig. 150). In
one aquatic genus (Maruina), the larvae have ventral
suckerlike disks. Aquatic larvae occur among vegetation and debris in shallow lentic habitats and along
margins of streams, and are frequently associated with
organically polluted water. They even occur in sink and
floor drains. Larvae feed near the surface on microorganisms and detritus. Life histories of aquatic species
are not well known; most are probably multivoltine,
especially in warm-water habitats. Larvae cannot be
identified to species.
k. Ptychopteridae (3 Genera, 16 Species) Larvae of
“phantom crane flies” are easily recognized by their
elongate caudal respiratory siphon and the presence of
pairs of prolegs on the first three abdominal segments
(Fig. 155). They live in very shallow water along margins of lentic or lotic habitats where there is an accumulation of detritus. Here, they obtain oxygen at the
surface through their caudal respiratory siphon and
probably feed on detritus and associated microorganisms. Their life history is poorly known because they
are generlly uncommon.
l. Simuliidae (11 Genera, 143 Species) “Blackfly”
larvae are uniquely shaped, having a swollen abdomen
that they attach to the substrate with a caudal sucker
(Fig. 156). They also have a relatively large head, and a
single ventral proleg on the thorax. They inhabit a
wide variety of lotic habitats where they are often
abundant on rocks, submerged wood, or vegetation in
fast to slow currents, each species usually having rather
specific ecological requirements. Most larvae are filter
feeders, using labral fans to filter diatoms and other
food from the current. Larvae without labral fans feed
by grazing on the substrate around them. Many species
are univoltine, while others are bivoltine or multivoltine. Species overwinter as either eggs or larvae, with
larvae of some species growing significantly during the
winter months. Unlike most other Nematocera, which
have only four larval instars, simuliid larvae have six
or sometimes seven instars. Pupation occurs at the site
of larval attachment, with pupae having a pair of
highly branched thoracic gills that aid in respiration.
Females usually require a blood meal for maturation of
eggs, and various species may be serious pests, biting
humans, livestock, and poultry. They also transmit
pathogenic organisms such as filarial nematodes, various viruses, and avian protozoan parasites.
m. Tanyderidae (2 Genera, 4 Species) The elongate
larvae of “primitive crane flies”, which may reach a
17. Diversity and Classification of Insects and Collembola
length of 18 mm, are rare in eastern and western North
America and have not been found in central North
America. They resemble chironomid larvae, but lack
anterior prolegs and have three pairs of long caudal filaments (Fig. 157). Larvae inhabit the shallow sand, silt,
or gravel margins of large streams, but, because they
are rare, their life history and feeding habits remain virtually unknown.
n. Thaumaleidae (2 Genera, 8 Species) The very
rare “solitary midge” larvae resemble chironomid larvae and reach a length of 12 mm, but have a single,
broad anterior and posterior proleg instead of paired
prolegs (Fig. 159). They also have dorsally sclerotized
segments and a pair of short, stalked spiracles dorsolaterally on the prothorax. Larvae inhabit vertical surfaces of rocks in cold, shady, mountain streams, where
the water film is thin enough so as not to cover them
completely. Here, they feed mostly on diatoms. Almost
nothing is known about their life history.
o. Tipulidae (20 Aquatic Genera, about 240 Species)
Larvae of “crane flies” differ from those of other Nematocera by having the posterior portion of the head
capsule incompletely sclerotized (Figs. 136 and 137)
and retracted into the thorax. In this very large family,
the vast majority of species are not aquatic, but there
are several genera with species that have aquatic larvae,
and these species are often widespread and abundant.
Because larvae of fewer than 10% of the species have
been described, it is difficult to know which species
have aquatic larvae. While most aquatic larvae are
found in lotic habitats, larvae in a few genera inhabit
shallow lentic habitats. Larvae of most aquatic species
have functional caudal spiracles that they may utilize to
obtain oxygen at the water surface, but those that live
in well-aerated streams probably obtain most of their
oxygen from the water through their cuticle. Larvae in
some genera are very large. Feeding habits vary widely,
with omnivores, herbivores, detritivores and carnivores
all represented. Most species are univoltine or bivoltine, but some that live in very cold water may be semivoltine. Pupation most often occurs in riparian habitats
adjacent to the larval habitat, but some species pupate
in shallow water and have thoracic respiratory horns
through which they breathe. Adults are short-lived.
The biology of Tipulidae was reviewed by Pritchard
(1983).
697
structures are greatly reduced and not rounded and
sclerotized as in Nematocera (orthorrhaphous Brachycera). Only in Stratiomyidae is there any significant head
structure, but heads of Stratiomyidae larvae are angulate (Fig. 138), partially retracted into the thorax, and
quite unlike heads of Nematocera. Mandibles of
Brachycera larvae move vertically, not horizontally as
in Nematocera. They lack subapical teeth and are usually referred to as “mouth hooks.”
The majority of species in all families, except Athericidae and Sciomyzidae, have terrestrial or semiaquatic
larvae, and it usually is not possible to know which
species have aquatic larvae because larvae of most
species of Brachycera remain unknown. Furthermore,
many genera that are known to have aquatic larvae,
also have terrestrial or semiaquatic larvae, so all species
within a genus cannot be assumed to be aquatic because larvae of one or more species have been found to
be aquatic. Because most larvae cannot be identified,
life histories are poorly known.
a. Athericidae (2 Genera, 4 Species) Aquatic larvae
of Atherix are distinctive, with pairs of ventral abdominal prolegs on the first seven abdominal segments and
with a single ventral proleg and a pair of fringed caudal projections on the eighth segment (Fig. 144). Another genus (Suragina) occurs in Texas; larvae were described by Webb (1994) and are similar to those of
Atherix. Atherix larvae commonly occur in riffles or
among vegetation in streams where they feed mostly on
insect larvae. They pupate in moist soil along the edge
of the stream. Adults are known to drink water and
perhaps feed on nectar; adult females of Suragina suck
blood from humans and cattle. Eggs of Atherix are laid
on vegetation or supports above the stream, with several females often contributing eggs to form a single,
large mass. Larvae drop into the stream upon hatching.
A 1-year life cycle is likely for most species.
2. Families of Aquatic Diptera — Suborder
Brachycera
b. Dolicopodidae (about 8 Aquatic Genera) The
“long-legged flies” are mostly terrestrial, but larvae of
a few species in eight genera are known to develop in a
wide variety of lotic and lentic habitats. The taxonomy
of larvae is so poorly known that identification to
genus is not reliable and it is not possible to clearly define which species are aquatic. Aquatic larvae differ
from other Brachycera by having a posterior spiracular
pit that is surrounded by four lobes (Fig. 143). They
are predaceous, feeding mostly on other insect larvae.
Pupae differ from other orthorrhaphous Brachycera by
having thoracic respiratory horns.
In larvae of Brachycera, either the head and external
head structures are replaced by an internal cephalopharyngeal skeleton (cyclorrhaphous Brachycera), or head
c. Empididae (about 8 Aquatic Genera) The small
(7 mm) larvae of “dance flies” are too poorly known
698
W. L. Hilsenhoff
to estimate the number of aquatic species. Larvae of
most species are terrestrial or semiaquatic. Most
aquatic larvae can be recognized because they have
seven or eight pairs of abdominal prolegs and a pair of
caudal respiratory appendages (Fig. 145), which are
not elongate and fringed as in Athericidae. Larvae are
predaceous on smaller animals, and most aquatic larvae live in rapid water of streams. Because larval taxonomy is so poorly known, almost nothing is known
about the life history of aquatic species.
d. Ephydridae (68 Genera) Species in most genera
have semiaquatic larvae. Some genera, however, have
aquatic species with larvae that live in vegetation or detritus in shallow habitats near shore, in algal mats, or
in unusual habitats such as hot springs and pools, and
ponds containing oil, as well as saline and alkaline
marshes, ponds and lakes, where larvae dominate the
insect fauna. Larvae of other species are terrestrial or
mine leaves of aquatic plants. “Shore fly” larvae are
less than 12 mm long and usually have a pair of very
short to elongate caudal respiratory tubes that frequently have a black sclerotized tip (Fig. 140). Many
also have abdominal prolegs. Most larvae feed on bluegreen and other algae, others feed on detritus, some are
predators. Aquatic species pupate in a puparium that
usually floats on the water surface. Eggs are laid singly
at the surface of the aquatic habitat. Although the family has received a great deal of attention, larval taxonomy is still poorly known and identification of larvae,
even at the generic level, is difficult or impossible.
Foote (1995) reviewed their biology.
e. Muscidae (about 8 Aquatic Genera) In this extremely large terrestrial family, larvae of several
aquatic species live in streams, marshes, or ponds; they
are mostly predators. Larvae are poorly known, but
most have a pair of very short caudal respiratory
tubes, a pair of ventral prolegs on the last abdominal
segment that are longer than the respiratory tubes, and
often creeping welts ventrally on other abdominal segments (Fig. 146). Eggs are laid in algae or debris, and
larvae usually remain near the surface. Pupation is in a
puparium, which remains at or near the surface. A
pair of respiratory siphons that are thrust through the
fourth segment of the puparium penetrate the water
surface.
f. Sciomyzidae (19 Genera, 175 Species) “Marsh
fly” larvae are predators or parasitoids of snails, slugs,
or fingernail clams, and only those species that attack
land snails or slugs are not aquatic. They generally
have a very wrinkled appearance, with girdles of
pseudopods on abdominal segments. The abdomen ter-
minates in a spiracular disk surrounded by eight to ten
lobes and contains a pair of spiracles, each surrounded
by numerous palmate hairs (Fig. 141). Larvae are
found in lentic habitats and margins of lotic habitats
inhabited by their prey. Here, they usually remain near
the surface with their caudal spiracles at the surface.
They swallow a large air bubble to keep themselves
afloat, and are usually buoyant enough to keep their
prey at the surface. Larvae pupate in a puparium that
either remains in the snail shell or floats at the surface.
Eggs are laid on emergent vegetation; newly hatched
larvae drop into the water and actively search for prey.
Adults of parasitoids oviposit directly on the shell of a
host snail. Species may be univoltine or multivoltine.
Because snails are intermediate hosts of liver flukes that
infect cattle and of schistosomes that attack humans,
sciomyzid larvae have been employed as a biological
control for snails. For this reason they have been extensively studied and larvae in several genera can be identified to species. Berg and Knudson (1978) reviewed the
biology and systematics.
g. Stratiomyidae (about 10 Aquatic Genera) Although most “soldier fly” larvae are terrestrial or semiaquatic, those of many species inhabit shallow, vegetated, lentic habitats. While often quite abundant,
larvae are so poorly known that identification at even
the generic level is not always reliable. The relatively
flat larvae are easily recognized by their mostly visible
truncate head (Fig. 138), caudal spiracles surrounded
by long hydrofuge hairs, and calcium carbonate crystals that cover their integument. Larvae hang suspended in the surface film by hydrofuge hairs that surround the spiracular disk, and use their palps to move
about. They may at times leave the water and inhabit
moist shoreline debris or plants. They feed on detritus
and also algae, differing from larvae of other orthorrhaphous Brachycera, which are predators. Pupation is
in a puparium, which may float in the aquatic habitat.
Adults feed mostly on flowers. Eggs are laid in masses,
either on submerged or emergent aquatic plants, or on
other objects in the water.
h. Syrphidae (about 7 Aquatic Genera) Most “rattailed maggosts” become fairly large and differ from
other Brachycera by being broad and blunt anteriorly.
Aquatic larvae are easily recognized by their very long,
extensile, caudal breathing tube (siphon) (Fig. 139).
Most also have ventral prolegs. Larvae inhabit shallow
lentic habitats or margins of lotic habitats, especially
areas high in decomposing organic matter. Here, they
feed on detritus and microorganisms. Larvae of some
species occur in tree holes. Because they obtain oxygen
from the air through their long respiratory siphon, they
17. Diversity and Classification of Insects and Collembola
frequently inhabit highly polluted areas such as sewage
lagoons. Larvae pupate in a puparium, with the puparium of some species having anterior respiratory horns.
Life histories are poorly known, but most species probably have more than one generation per year. Adults
are called “flower flies” because they feed on nectar
from flowers. Eggs are laid in masses on the debris or
vegetation that the larvae inhabit.
i. Tabanidae (about 4 Aquatic Genera) Most “deerfly” or “horsefly” larvae develop in wet soil that may
be closely associated with aquatic habitats. Some larvae in four genera are known to be aquatic, and a few
in other genera also may be aquatic. Many North
American species have been reared and described, and
larval keys have been developed, but identification is
still risky because larvae of some species remain unknown. Larvae are recognized by the absence of distinct prolegs and the presence of a girdle of six or more
pseudopods on most abdominal segments (Fig. 142).
Aquatic larvae have been collected from stream riffles,
shallow stream margins, and shallow vegetated lentic
habitats. Most species are probably univoltine, but bivoltine and semivoltine species likely occur. Larvae are
predators on other aquatic macroinvertebrates; pupation of aquatic species takes place in semiaquatic areas.
Most adult females feed on blood of humans, livestock, and other animals, and are often a persistent
nuisance. Eggs are laid in masses on vegetation above
the larval habitat.
IV. SEMIAQUATIC COLLEMBOLA — SPRINGTAILS
Until recently Collembola were regarded as insects, but
most people no longer consider them to be in the class
Insecta and place them in the class Entognatha or in a
subclass of Hexopoda. The order Collembola is only
partially aquatic, with the vast majority of the nearly
700 North American species inhabiting moist terrestrial habitats. Species that are collected from aquatic
habitats are usually found on the water surface and are
semiaquatic. Waltz and McCafferty (1979) reported
that 96 species of springtails had been collected at least
once from freshwater or coastal marine habitats in
North America, but most of these records were believed to be incidental occurrences of terrestrial species.
They regarded 10 species as semiaquatic and five as riparian with a propensity to venture onto the water surface. These 15 species exhibited various degrees of
adaptation for the semiaquatic environment and are
discussed in detail by Waltz and McCafferty, who also
key species most likely to be found in freshwater habitats. Keys and descriptions for all North American
699
species of Collembola are provided by Christiansen and
Bellinger (1980).
Collembola are small, wingless arthropods, usually
much less than 6 mm long. They resemble insects by
having a distinct head with one pair of segmented antennae, a three-segmented thorax with three pairs of
legs, and a segmented abdomen without legs (Figs. 160
and 161).
They differ from insects in having only six abdominal segments and in having their maxillae and
mandibles concealed by the wall of the head capsule
with which their labium is fused. They also have the
tibia and tarsus of each leg fused into a single segment.
Except for their smaller size and lack of genitalia,
young Collembola closely resemble adults.
Springtails can be readily recognized by a cylindrical collophore or ventral tube (Fig. 160 and 161) on
the first abdominal segment and by the furcula, a midventral appendage on the fourth abdominal segment
(Fig. 160 and 161) that is present in most species and
in all that are known to be semiaquatic. The furcula
consists of a basal piece, the manubrium, and two arms
(Fig. 162). The basal segment of each arm is the dens
and the apical segment is the mucro. The furcula can
be folded forward under the abdomen where it is held
in place by the tenaculum, a midventral clasplike structure on the third abdominal segment. When the furcula
is released, it springs backward and propels the
Collembola several centimeters through the air, hence
the common name “springtail.” In most semiaquati
species there is a widening of each mucro or lamellae
on each mucro. This permits greater contact with the
surface film, allowing individuals to spring farther
when on the water surface.
The ecology and biology of Collembola remains
very poorly known. Semiaquatic species are most often
associated with lentic freshwater habitats; none occur
on the ocean and reports from lotic habitats are rare.
They feed primarily on algae, detritus, and other organic material in the surface film; some may eat bacteria. Adults do not copulate like insects. Instead males
deposit stalked or sessile spermatophores, which are
gathered by the females; usually no contact between
sexes is involved. Eggs are laid in moist habitats; in at
least one species they are laid beneath the water surface
and newly hatched young are truly aquatic, but once
they pass through the surface film and come in contact
with the air they remain semiaquatic. Eggs usually
hatch within a few weeks, except those that diapause
over the winter or during periods of dryness. The young
reach sexual maturity within a few weeks and molt
from three to seven times. In some species the young
may enter into diapause. Most adults live a few weeks
to a few months, and unlike insects, they continue to
700
W. L. Hilsenhoff
molt throughout their adult life, with more than fifty
molts reported for one species. Some species are coldhardy and may be active at temperatures near freezing;
however, most species overwinter as diapausing eggs.
Because of their springing ability Collembola are
often difficult to capture. Semiaquatic species may be
collected most easily by forcing them to jump into a
white pan containing either 95% alcohol with 3%
glacial acetic acid, or water with detergent to prevent
them from jumping from the pan. Floating sticky traps
may also be used to capture them. Christiansen and
Bellinger (1980) recommend 70 – 95% isopropanol or
80 – 95% ethanol for preservation, with 1 – 2% glycerine added to prevent accidental desiccation. A taxo-
nomic key to families of semiaquatic Collembola appears in Section V.J.
V. IDENTIFICATION OF THE FRESHWATER
INSECTS AND COLLEMBOLA
The following taxonomic keys are separated by class
(Entognatha and Insecta) and orders (Insecta only).
Collembola, which usually are much less than 6 mm
long, can be distinguished from small larval insects by
the possession ventrally of a collophore (ventral tube)
on the first abdominal segment and a furcula on the
fourth (Figs. 158 and 159).
A. Taxonomic Key to Orders of Freshwater Insects
1a.
Thorax with 3 pairs of segmented legs ...............................................................................................................................................3
1b.
Thorax without segmented legs ..........................................................................................................................................................2
2a(1b).
Mummylike, with developing adult structures; in a case, often silk-cemented and contains vegetable or mineral matter......................
.................................................................................................................................................................................pupae (not keyed)
2b.
Not in a case; mobile larvae, mostly with prolegs, pseudopods, or creeping welts on one or more segments (Figs. 145 – 147 and
152 – 159).......................................................................................................................................................................Diptera larvae
3a(1a).
With large functional wings, which may be shelllike, or leatherlike at the base ..................................................................................4
3b.
Wingless, or with developing wings (wing-pads) or brachypterous wings...........................................................................................6
4(3a).
All wings completely membranous, with numerous veins ..................................................ovipositing or terrestrial adults (not keyed)
4b.
Mesothoracic wings hardened and shell-like (Figs. 108, 117, and 118), or leatherlike in basal half (Figs. 87 and 93 – 95) .................5
5a(4b).
Chewing mouthparts; mesothoracic wings hard, shell-like (Figs. 108, 117, 118) ......................................................Coleoptera adults
5b.
Sucking mouthparts formed into a broad or narrow tube (Figs. 88 and 90); mesothoracic wings hardened in basal half (Fig. 87);
.......................................................................................................................................................Heteroptera ( Hemiptera) adults
6a(3b).
With 2 or 3 long, filamentous terminal appendages (Figs. 9, 29, 45, 48, 49, 54, 56, and 58) .............................................................7
6b.
Terminal appendages absent, or with only 1 or 2 segments ................................................................................................................8
7a(6a).
Sides of abdomen with featherlike, platelike, or leaflike gills (Figs. 4, 11 – 15, and 25 – 30); usually with 3 tail filaments, occasionally
only 2; tarsi with 1 claw.....................................................................................................................................Ephemeroptera larvae
7b.
Gills absent from at least middle abdominal segments (Figs. 45, 48, 49, 54, 56, and 58); 2 tail filaments; tarsi with 2 claws ...............
Plecoptera larvae
8a(6b).
Labium formed into an elbowed, extensile grasping organ (Figs. 32 – 34); abdomen terminating in 3 lamellae (Fig. 31) or 5 triangular points (Fig. 44) .......................................................................................................................................................Odonata larvae
8b.
With sucking or chewing mouthparts; abdomen not terminating in 3 lamellae or 5 triangular points ................................................9
9a(8b).
Mouthparts sucking, formed into a broad or narrow tube (Figs. 88 and 90) or a pair of long stylets (Fig. 102)...............................10
9b.
Mouthparts not sucking, not formed into a tube or pair of stylets ...................................................................................................11
10a(9a).
Parasitic on sponges; mouthparts a pair of long stylets (Fig. 102); all tarsi with 1 claw (Fig. 102) ...........Neuroptera; Sisyridae larvae
10b.
Free-living; mouthparts a broad or narrow tube (Figs. 88 and 90); mesotarsi with 2 claws (Figs. 87 and 93 – 96)................................
......................................................................................................................................Heteroptera ( Hemiptera) adults or nymphs
11a(9b)
Ventral prolegs on abdominal segments 3 – 6, each with a ring of fine hooks (Fig. 103) ..........................Lepidoptera; Pyralidae larvae
11b.
Abdomen without ventral prolegs on abdominal segments 3 – 6 that have a ring of hooks ...............................................................12
12a(11b).
Antennae extremely small, inconspicuous, one-segmented (Figs. 63, 78, and 80).....................................................Trichoptera larvae
12b.
Antennae elongate, with 3 or more segments ...................................................................................................................................13
17. Diversity and Classification of Insects and Collembola
701
13a(12b).
Without long lateral filaments (Figs. 120 – 130) ........................................................................................................Coleoptera larvae
13b.
With long lateral filaments (Figs. 85, 86, and 119) ...........................................................................................................................14
14a(13b).
A single claw on each tarsus (Fig. 121 – 124) or abdomen terminating in 2 slender filaments or a median proleg with 4 hooks (Fig.
119) ..........................................................................................................................................................................Coleoptera larvae
14b.
Each tarsus with 2 claws; abdomen terminating in a single slender filament (Fig. 85) or in 2 prolegs, each with 2 hooks (Fig. 86)
..............................................................................................................................................................................Megaloptera larvae
B. Taxonomic Key to Families of Freshwater Ephemeroptera Larvae
1a.
Mandibles with large forward-projecting tusks (Fig. 3); gills on abdominal segments 2 – 7 with fringed margins (Fig. 4) and projecting laterally or dorsally over abdomen ...............................................................................................................................................2
1b.
Mandibles without large tusks; conspicuous fringed gills absent or projecting ventrolaterally............................................................4
2a(1a).
Gills dorsal, curving up over abdomen; protibiae fossorial (Fig. 5).....................................................................................................3
2b.
Gills lateral, projecting from sides of abdomen; protibiae slender, subcylindrical (Fig. 6); East, Midwest......................Potamanthidae
3a(2a).
Apex of metatibiae rounded (Fig. 7); mandibular tusks curved inward and downward apically ..................................Polymitarcyidae
3b.
Apex of metatibiae projected into an acute point ventrally (Fig. 8); mandibular tusks curved upward or outward apically
........................................................................................................................................................................................Ephemeridae
4a(1b).
Mesonotum modified into a carapace-like structure that covers the gills on abdominal segments 1 – 6 (Fig. 9) ...................Baetiscidae
4b.
Mesonotum not modified into a carapace, gills exposed.....................................................................................................................5
5a(4b).
Head and pronotum with pads of long, dark setae on each side (Fig. 10); fringed gills projecting ventrolaterally; Southeast WI
.........................................................................................................................................................................................Behningiidae
5b.
Head and pronotum without pads of long, dark setae; gills not fringed and projecting ventrolaterally ..............................................6
6a(5b).
Gills on abdominal segment 2 operculate or semi-operculate, covering or partially covering gills on succeeding segments (Figs.
11 – 14)...............................................................................................................................................................................................7
6b.
Gills on abdominal segment 2 similar to those on succeeding segments or absent ..............................................................................9
7a(6a).
Operculate gills oval or subtriangular and well-separated from each other mesally (Figs. 11 and 12); gills on segments 3 – 6 without
fringed margins .............................................................................................................................................................Leptohyphidae
7b.
Operculate gills quadrate and meeting along mesal edge (Figs. 13 and 14); gills on segments 3 – 6 with fringed margins ...................8
8a(7b).
Operculate gills fused mesally (Fig. 13); East..............................................................................................................Neoephemeridae
8b.
Operculate gills not fused mesally, but overlapping (Fig. 14) .................................................................................................Caenidae
9a(6b).
Gills absent from abdominal segment 2 and sometimes also from 1 and 3, gills on segments 3 or 4 may be operculate (Fig. 15)
.....................................................................................................................................................................................Ephemerellidae
9b.
Gills present on abdominal segments 1 or 2 to 7 ..............................................................................................................................10
10a(9b).
Head flattened dorsoventrally; eyes and antennae dorsal (Fig. 16); gills a single lamella, often with a fibrilliform tuft .....................11
10b.
Head and body not dorsoventrally flattened; eyes, and usually also antennae, along lateral margin of head (Fig. 17) ......................13
11a(10a).
Distal maxillary palpomere at least 4 times as long as galea-lacinia and visible from above (Fig. 18); Northeast............Arthropleidae
11b.
Distal maxillary palpomere much shorter, usually not visible from above ........................................................................................12
12a(11b).
Gills with a ventral fingerlike projection on lamellae (Fig. 19); tarsal claws very long: West, Midwest, Southeast ..........Pseudironidae
12b.
Gill lamellae without such a projection; claws not exceptionally long............................................................................Heptageniidae
13a(10b).
Protarsal claws much shorter than tibiae and those on other tarsi (Figs. 20 and 22); claws on meso- and metatarsi long and slender,
about as long as tibiae (Fig. 21)........................................................................................................................................................14
13b.
Claws on all tarsi similar in structure and length..............................................................................................................................15
14a(13a).
Protarsal claws bifid (Fig. 20); no spinose pad on procoxae; Canada, midwest and, southeast United States ..............Metretopodidae
14b.
Protarsal claws simple, with long, slender denticles (Fig. 22); spinose pad present on procoxae; West, Upper MI........Ametropodidae
15a(13b).
Prothoracic legs with dense row of long setae along inner surface (Fig. 23)......................................................................................16
15b.
Prothoracic legs without dense row of setae along inner surface ......................................................................................................17
702
W. L. Hilsenhoff
FIGURE 3 Ephemeridae; head of Pentaqenia (dorsal view) showing mandibular tusk (T). FIGURE 4
Ephemeridae; gills of Hexaqenia. FIGURE 5 Ephemeridae; prothoracic leg of Hexaqenia. FIGURE 6
Potamanthidae; prothoracic leg of Anthopotamus. FIGURE 7 Polymitarcyidae; metatibia and tarsus of
Ephoron. FIGURE 8 Ephemeridae; metatibia and tarsus of Ephemera. FIGURE 9 Baetiscidae; Baetisca
(dorsal view). FIGURE 10 Behningiidae; head and pronotum of Dolania. FIGURE 11 Leptohyphidae;
abdomen of Tricorythodes (dorsal view) showing operculate gill (OG). FIGURE 12 Leptohyphidae; abdomen
of Leptohyphes (dorsal view) showing operculate gill (OG). FIGURE 13 Neoephemeridae; abdomen of
Neoephemera (dorsal view) showing operculate gill (OG). FIGURE 14 Caenidae; abdomen of Caenis (dorsal
view) showing operculate gill (OG).
16a(15a).
Gills on abdominal segment 1 dorsolateral; gills lamellate, tapered apically, with a basal fibrilliform tuft; gill tuft at base of procoxae (Fig. 23) ..................................................................................................................................................................Isonychiidae
16b.
Gills on abdominal segment 1 ventral; gills on abdominal segments 2 – 7 small, either lanceolate with a posterior fringe (Fig. 24), or
rounded and platelike; no gill tuft at base of procoxae; West, midwest and and southeast United States.......................Oligoneuriidae
17a(15b).
Gills forked (Fig. 25), or bilamellate and terminating in a point (Fig. 26), or terminating in filaments (Fig. 27) ..........Leptophlebiidae
17b.
Gills single or double lamellae (Figs. 28 and 29) ..............................................................................................................................18
17. Diversity and Classification of Insects and Collembola
703
FIGURE 15 Ephemerellidae; abdomen of Eurylophella (dorsal view) showing operculate gill (OG). FIGURE
16 Heptageniidae; head of Stenonema (dorsal view). FIGURE 17 Siphlonuridae; head of Siphlonurus
(dorsal view). FIGURE 18 Arthropleidae; head of Arthroplea (dorsal view) showing maxillary palp (MP).
FIGURE 19 Pseudironidae; gill lamella of Pseudiron (ventral view). FIGURE 20 Metretopodidae; protarsus
and tibia of Siphloplecton. FIGURE 21 Metretopodidae; metatarsus and tibia of Siphloplecton. FIGURE 22
Ametropodidae; protarsus and tibia of Ametropus. FIGURE 23 Isonychiidae; prothoracic leg of Isonychia
showing gill (G). FIGURE 24 Oligoneuriidae; gill on abdominal segment 3 of Homoeoneuria. FIGURE 25
Leptophlebiidae; gill on abdominal segment 3 of Paraleptophlebia. FIGURE 26 Leptophlebiidae; gills on
abdominal segment 3 of Leptophlebia. FIGURE 27 Leptophlebiidae; gill on abdominal segment 3 of
Habrophlebia. FIGURE 28 Siphlonuridae; gills on abdominal segment 2 of Siphlonurus. FIGURE 29
Baetidae; Baetis (dorsal view). FIGURE 30 Ameletidae; gill on abdominal segment 3 of Ameletus.
18a(17b).
Tibiae and tarsi bowed; claws very long and slender; metatarsal claws about as long as tarsi; Illinois, Wisconsin, South Carolina,
and Georgia .......................................................................................................................................................Acanthametropodidae
18b.
Tibiae and tarsi not bowed; claws not as above................................................................................................................................19
19a(18b).
Abdominal segments 8 and 9 produced posterolaterally into distinct, flattened spines (Figs. 13 – 15); if spines are weak, antenna
more than twice width of head .........................................................................................................................................................20
704
W. L. Hilsenhoff
19b.
Abdominal segments 8 and 9 without such spines (Fig. 29), if weak spines are present, antenna less than twice width of head
...............................................................................................................................................................................................Baetidae
20a(19a).
Gills single, with sclerotized band on ventral margin and little or no tracheation (Fig. 30); maxillae with crown of pectinate spines
...........................................................................................................................................................................................Ameletidae
20b.
Gills with well-developed tracheation (Fig. 28); maxillae without a crown of pectinate spines .......................................Siphlonuridae
C. Taxonomic Key to Families of Freshwater Odonata Larvae
1a.
Abdomen terminating in 3 caudal lamellae, longest more than half length of abdomen (Fig. 31) .......................suborder Zygoptera 2
1b.
Abdomen terminating in 3 stiff, pointed valves and 2 pointed cerci, longest 1 / 3 length of abdomen (Fig. 44) ..................................
..........................................................................................................................................................................suborder Anisoptera 5
2a(1a).
Zygoptera: first antennal segment as long as, or longer than, remaining segments combined (Fig. 31); prementum with deep, median
cleft (Fig. 32) ................................................................................................................................................................Calopterygidae
2b.
First antennal segment much shorter than others combined; prementum with at most a very small median cleft (Figs. 33 and 34) ....3
3a(2b).
Basal half of prementum greatly narrowed and elongate (Fig. 33); prementum in repose extends back to or past mesocoxae .Lestidae
3b.
Basal half of prementum not greatly narrowed (Fig. 34); prementum in repose extends only to procoxae..........................................4
4a(3b).
Caudal lamellae divided into a thick basal half and thin, lighter colored distal half (Fig. 35); one dorsal seta on each side of prementum; Texas......................................................................................................................................................................Protoneuridae
4b.
Caudal lamellae not distinctly divided as above; usually with 2 or more dorsal setae on each side of prementum .......Coenagrionidae
5a(1b).
Anisoptera: Prementum flat or nearly so, without dorsal setae; palpal lobes also flat and usually without stout setae ........................6
5b.
Prementum rounded, spoon-shaped, and usually with dorsal setae; palpal lobes also rounded, always with stout setae, and covering
face to base of antennae .....................................................................................................................................................................8
6a(5a).
Antennae 4-segmented (Fig. 36); pro- and mesotarsi 2-segmented......................................................................................Gomphidae
6b.
Antennae 6- or 7-segmented (Figs. 37 and 38); pro- and mesotarsi 3-segmented ................................................................................7
7a(6b).
Antennal segments slender, not hairy (Fig. 37) .....................................................................................................................Aeshnidae
7b.
Antennal segments short, thick, and hairy (Fig. 38); mountain seeps ..................................................................................Petaluridae
8a(5b).
Distal margin of palpal lobes with large, irregular teeth (Fig. 39) ..............................................................................Cordulegastridae
8b.
Distal margin of palpal lobes with small, even crenulations (Figs. 40 and 41) or nearly straight (Fig. 42) ..........................................9
9a(8b).
Head with a prominent, almost erect, thick frontal process between bases of antennae (Fig. 43); legs very long, apex of each metafemur reaching to or beyond apex of abdominal segment 8; metasternum with broad, median tubercle ............................Macromiidae
9b.
Head without a prominent frontal process (except in Neurocordulia molesta); legs shorter, apex of metafemurs usually not reaching
apex of abdominal segment 8; metasternum without a median tubercle ...........................................................................................10
10a(9b).
Crenulations on palpal lobes very shallow, 1 / 6 to 1 / 10 as long as wide (Fig. 42), or separated by minute notches bearing setae
..................................................................................................................................................................................most Libellulidae
10b.
Crenulations on palpal lobes large, at least 1 / 4 as long as wide (Figs. 40 and 41)............................................................................11
11(10b).
Lateral spines on abdominal segment 8 as long or longer than middorsal length of segment 9 (Fig. 44).............................Libellulidae
Lateral spines on abdominal segment 8 shorter than middorsal length of segment 9..........................................................Corduliidae
D. Taxonomic Key to Families of Freshwater Plecoptera Larvae
1a.
Conspicuous finely branched gills present ventrally or laterally on all thoracic segments (Fig. 2) .......................................................2
1b.
Gills absent, confined to prosternum, or unbranched .........................................................................................................................3
2a(1a).
Finely branched gills on abdominal sterna 1, 2 and sometimes 3 ..................................................................................Pteronarcyidae
2b.
Gills absent from first 3 visible abdominal sterna.....................................................................................................................Perlidae
3a(1b).
Thoracic sterna produced into plates that overlap succeeding segment (Fig. 45); single, double, or forked gills behind meso- and
metacoxae; form roach-like; mountain streams ................................................................................................................Peltoperlidae
17. Diversity and Classification of Insects and Collembola
705
FIGURE 31 Calopterygidae; Calopteryx (dorsal view). FIGURE 32 Calopterygidae; prementum of Hetaerina (dorsal view). FIGURE 33 Lestidae; prementum of Lestes (dorsal view). FIGURE 34 Coenagrionidae; prementum of Enallagma (dorsal view) showing dorsal setae (DS), palpal lobe (PL), and palpal setae
(PS). FIGURE 35 Protoneuridae; caudal lamella of Protoneura (lateral view). FIGURE 36 Gomphidae;
antenna of Gomphus. FIGURE 37 Aeshnidae; antenna of Anax. FIGURE 38 Petaluridae; right antenna
of Tachopteryx (dorsal view). FIGURE 39 Cordulegastridae; palpal lobe of Cordulegaster (dorsal view).
FIGURE 40 Corduliidae; palpal lobe of Neurocordulia (dorsal view). FIGURE 41 Corduliidae; palpal
lobe of Cordulia (dorsal view). FIGURE 42 Libellulidae; palpal lobe of Tramea (dorsal view). FIGURE 43
Macromiidae; head of Macromia (dorsal view). FIGURE 44 Libellulidae; abdominal segments 7 – 10 of
Pantala (dorsal view) showing epiproct (E), cerci (C), and paraprocts (P).
3b.
Thoracic sterna not produced into overlapping plates; coxal gills, if present, segmented and not apically pointed; form usually elongate, not roachlike..............................................................................................................................................................................4
4a(3b).
Tips of glossae situated much behind tips of paraglossae (Fig. 46) .....................................................................................................5
4b.
Tips of glossae produced nearly as far forward as tips of paraglossae (Fig. 47) ..................................................................................6
706
W. L. Hilsenhoff
FIGURE 45
Peltoperlidae: Peltoperla (ventral view) with legs removed beyond coxae. FIGURE 46
Chloroperlidae; labium of Alloperla (ventral view) showing glossae (G) and paraglossae (P). FIGURE 47
Nemouridae; labium of Nemoura (ventral view) showing glossae (G) and paraglossae (P). FIGURE 48
Perlodidae; Isoperla (dorsal view). FIGURE 49 Chloroperlidae; Haploperla (dorsal view).
5a(4a).
Cerci almost as long or longer than abdomen; metathoracic wing-pads with inner and outer margins strongly diverging from axis of
body (Fig. 48); head and thorax usually with a distinct pattern ...........................................................................................Perlodidae
5b.
Cerci distinctly shorter than abdomen; metathoracic wing-pads with inner and outer margins nearly parallel to axis of body (Fig.
49); head and thorax usually not patterned ..................................................................................................................Chloroperlidae
6a(4b).
Second tarsal segment (lateral view) about as long as, or longer than first (Fig. 50); single segmented gills on inner side of each coxa
(Fig. 51), or ninth abdominal sternum greatly produced (Fig. 52)..............................................................................Taeniopterygidae
6b.
Second tarsal segment much shorter than first (Fig. 53); gills absent from inner side of coxae and ninth sternum not produced ........7
7a(6b).
Robust larvae, with extended metathoracic legs reaching to or beyond tip of abdomen; metathoracic wing-pads with inner and
outer margins strongly diverging from axis of body (Fig. 54)............................................................................................Nemouridae
17. Diversity and Classification of Insects and Collembola
707
FIGURE 50
Taeniopterygidae; tarsal segments (1, 2, 3) of Taeniopteryx (lateral view). FIGURE 51
Taeniopterygidae; mesosternum of Taeniopteryx showing gills (G) at base of coxae. FIGURE 52
Taeniopterygidae; ninth abdominal sternum (9) of Strophopteryx (ventral view). FIGURE 53 Nemouridae;
tarsal segments of Nemoura (lateral view). FIGURE 54 Nemouridae; Amphinemura (dorsal view). FIGURE
55 Capniidae; abdomen of Allocapnia (lateral view) showing lateral fold (LF). FIGURE 56 Capniidae;
Paracapnia (dorsal view). FIGURE 57 Leuctridae; abdomen of Leuctra (lateral view) showing lateral fold
(LF). FIGURE 58 Leuctridae; Leuctra (dorsal view).
7b.
Elongate larvae, with extended metathoracic legs reaching well short of tip of abdomen; metathoracic wing-pads with inner and
outer margins nearly parallel to axis of body (Figs. 56, and 58) .........................................................................................................8
8a(7b).
Abdominal segments 1 – 9 divided by a membranous ventrolateral fold (Fig. 55); metathoracic wing-pads, if present, about same
distance apart as mesothoracic wing-pads and often short and broad (Fig. 56) ....................................................................Capniidae
8b.
Abdominal segments 7 – 9 not divided by a membranous fold (Fig. 57); metathoracic wing-pads similar in shape to mesothoracic
wing-pads, but distinctly closer together (Fig. 58) ................................................................................................................Leuctridae
708
W. L. Hilsenhoff
E. Taxonomic Key to Families of Freshwater Trichoptera Larvae
1a.
Larvae in spiral case of sand grains or tiny stones that resembles a snail shell (Fig. 59); anal claws with many short teeth forming a
comb, not hook-shaped at apex (Fig. 60) ....................................................................................................................Helicopsychidae
1b.
Case not like a snail shell or absent; anal claws not forming a comb, and hook-shaped at apex.........................................................2
2a(1b).
Each thoracic segment covered with a single dorsal plate, which may have a mesal or transverse fracture line...................................3
2b.
Metanotum mostly membranous, having only scattered hairs or small plates, or with 2 or more sclerites..........................................5
3a(2a).
Abdomen with rows of branched gills ventrally; no portable case...............................................................................Hydropsychidae
3b.
Abdomen without branched gills ventrally; often with a portable case ...............................................................................................4
4a(3b).
Anal prolegs very short and with a stout claw; abdominal tergum 9 sclerotized; fifth instar larvae with abdomen much enlarged; in
a barrel- or purse like case (Figs. 61, and 62), or in a flat, silk case ................................................................................Hydroptilidae
4b.
Anal prolegs elongate with a projecting claw; abdominal tergum 9 entirely membranous; abdomen not enlarged; larvae in sand retreats; southwestern United States .......................................................................................................................................Ecnomidae
5a(2b).
Antennae long, at least 6 times as long as wide, and arising near base of mandibles (Fig. 63); or mesonotum membranous, except
for a pair of sclerotized, narrow, curved or angled bars (Fig. 64); larvae in a case ............................................................Leptoceridae
5b.
Antennae very short, not more than 3 times as long as wide, often inconspicuous and arising at various points; mesonotum never
with a pair of sclerotized, narrow, curved or angled bars; with or without case .................................................................................6
6a(5b).
Meso- and metanotum entirely membranous, or with only weak sclerites at SA-1 of mesonotum (Fig. 65); pronotum never with an
anterolateral projection; larvae with or without a case .......................................................................................................................7
6b.
Meso- and metanotum with some conspicuous sclerotized plates (Figs. 83 and 84); pronotum sometimes with an anterolateral projection; larvae in a case (Fig. 66) ......................................................................................................................................................15
7a(6a).
Abdominal segment 9 with dorsum entirely membranous; no portable case (Fig. 67).........................................................................8
7b.
Abdominal segment 9 bearing a sclerotized dorsal plate; with or without portable case...................................................................12
8a(7a).
Tibia and tarsus fused to form a single segment on all legs; mesopleura with a forward projecting lobe (Fig. 68); southwestern
United States ............................................................................................................................................................Xiphocentronidae
8b.
Tibia and tarsus distinct on all legs; mesopleura without a projecting lobe.........................................................................................9
9a(8b).
Protrochantins broad, hatchet-shaped (Fig. 69) .............................................................................................................Psychomyiidae
9b.
Protrochantins pointed or poorly developed.....................................................................................................................................10
10a(9b).
Protrochantins poorly developed; head without markings; labrum membranous and T-shaped (Fig. 70) .....................Philopotamidae
10b.
Protrochantins pointed anteriorly (Fig. 71); head usually with dark markings or dark or light muscle scars; labrum sclerotized and
widest near base ...............................................................................................................................................................................11
11a(10b).
Tarsi broad and densely pilose (Fig. 72); mandibles short and triangular, each with a large, thick mesal brush; East, Central..............
....................................................................................................................................................................................Dipseudopsidae
11b.
Tarsi with little or no pile and with a large claw (Fig. 67); mandibles elongate........................................................Polycentropodidae
12a(7b).
Prosternal horn present (Fig. 80); SA-3 on meso- and metanotum with a small sclerite and a cluster of setae (Fig. 65); case of vegetation, spirally wound or a series of rings and readily abandoned ......................................................................................Phryganeidae
12b.
Prosternal horn absent; SA-3 on meso- and metanotum without a sclerite and usually with a single seta; case, if present, of sand and
pebbles .............................................................................................................................................................................................13
13a(12b).
Anal claws very small, much shorter than elongate sclerite on anal legs (Fig. 73); larvae with turtlelike case of sand and pebbles,
which is readily abandoned........................................................................................................................................Glossosomatidae
13b.
Anal claws large, as long as elongate sclerite on anal legs (Fig. 74); larvae without a case................................................................14
141(13b). Profemurs with ventral projection to form chelate legs (Fig. 75); southwestern United States........................................Hydrobiosidae
14b.
Prothoracic legs not chelate..........................................................................................................................................Rhyacophilidae
15a(6b).
Claws of metathoracic legs very small or a slender filament, those of other legs long; case of sand, usually with lateral flanges ...........
..........................................................................................................................................................................................Molannidae
15b.
Claws of metathoracic legs as long as those of mesothoracic legs; case never of sand with lateral flanges ........................................16
16a(15b).
Pronotum divided by a sharp furrow across middle, area in front of furrow depressed (Fig. 76); no dorsal or lateral humps on
abdominal segment 1 ..................................................................................................................................................Brachycentridae
17. Diversity and Classification of Insects and Collembola
709
FIGURE 59 Helicopsychidae; case of Helicopsyche (dorsal view). FIGURE 60 Helicopsychidae; claw on
anal proleg of Helicopsyche. FIGURE 61 Hydroptilidae; case of Oxyethira (lateral view). FIGURE 62
Hydroptilidae; case of Hydroptila (lateral view). FIGURE 63 Leptoceridae; head of Oecetis (lateral view)
showing antenna (A). FIGURE 64 Leptoceridae; head and thoracic terga of Ceraclea. FIGURE 65
Phryganeidae; head and thoracic terga of Oligostomis showing location of setal areas (SA). FIGURE 66
Limnephilidae; Hesperophylax (lateral view). FIGURE 67 Polycentropodidae; Polycentropus (lateral view).
FIGURE 68 Xiphocentronidae; pro- and mesothorax of Xiphocentron (lateral view) showing projecting
lobe (L). FIGURE 69 Psychomyiidae; pronotum (P) and protrochantin (T) of Psychomyia. FIGURE 70
Philopotamidae; head of Chimarra (dorsal view) showing labrum (L). FIGURE 71 Polycentropodidae;
pronotum (P) and protrochantin (T) of Polycentropus. FIGURE 72 Dipseudopsidae; tarsus of
Phylocentropus.
16b.
Pronotum with at most a shallow furrow; lateral humps and usually dorsal hump present on abdominal segment 1.......................17
17a(16b).
Pronotum divided obliquely by a sharp carina that terminates as a projecting, rounded lobe anterolaterally (Fig. 77); East, local .......
..............................................................................................................................................................................................Beraeidae
17b.
Pronotum without a distinct carina terminating in a rounded anterolateral lobe..............................................................................18
710
W. L. Hilsenhoff
FIGURE 73 Glossosomatidae; last abdominal segment of Glossosoma (lateral view) showing anal proleg.
FIGURE 74 Rhyacophilidae; last abdominal segment of Rhyacophila (lateral view) showing anal proleg.
FIGURE 75 Hydrobiosidae; pronotum and prothoracic leg with chelate femur (F) of Atopsyche (posterolateral view). FIGURE 76 Brachycentridae; pronotum and head of Brachycentrus (lateral view). FIGURE 77
Beraeidae; pronotum of Beraea (dorsal view) showing carina (C). FIGURE 78 Lepidostomatidae; head of
Lepidostoma (dorsal view) showing antenna (A). FIGURE 79 Calamoceratidae; labrum of Heteroplectron
(dorsal view). FIGURE 80 Limnephilidae; head and prothorax of Platycentropus (lateral view) showing
antenna (A) and prosternal horn (H). FIGURE 81 Sericostomatidae; pronotum (P), protrochantin (T), and
coxa (C) of Aqarodes. FIGURE 82 Odontoceridae; pronotum (P), protrochantin (T), and coxa (C) of
Psilotreta. FIGURE 83 Goeridae; thoracic terga of Goera showing mesepisternum (M). FIGURE 84
Uenoidae; thoracic terga of Neophylax.
17. Diversity and Classification of Insects and Collembola
711
18a(17b).
Antennae located extremely close to eyes (Fig. 78); median dorsal hump absent from abdominal segment 1............Lepidostomatidae
18b.
Antennae midway between eyes and base of mandibles or near base of mandibles; dorsal hump present on abdominal segment
1.......................................................................................................................................................................................................19
19a(18b).
Anterolateral angles of pronotum produced and divergent; labrum with transverse dorsal row of about 18 long setae (Fig. 79); case
of vegetation; East and West ......................................................................................................................................Calamoceratidae
19b.
Anterolateral angles of pronotum if produced, not divergent; labrum without a dorsal row of about 18 long setae; case of various
materials ..........................................................................................................................................................................................20
20a(19b).
Prosternal horn absent; antennae close to base of mandibles ............................................................................................................21
20b.
Prosternal horn present (Fig. 80); antennae about midway between base of mandibles and eye (Fig. 80) .........................................22
21a(20a).
Protrochantin large, hook-shaped (Fig. 81); anal prolegs each with about 30 long setae............................................Sericostomatidae
21b.
Protrochantins small, not hook-shaped (Fig. 82); anal prolegs each with about 5 long setae.........................................Odontoceridae
22a(20b).
Mesepisternum formed anteriorly into a sharp, elongate process (Fig. 83) or a rounded spiny prominence............................Goeridae
22b,
Mesepisternum not enlarged anteriorly as above ..............................................................................................................................23
23a(22b).
Mesonotum with an anterior mesal notch (Fig. 84); SA-1 of metanotum unsclerotized and with only 1 or 2 setae (Fig. 84). ...............
..............................................................................................................................................................................................Uenoidae
23b.
Mesonotum without an anterior notch; SA-1 of metanotum with a sclerotized plate and / or more than 2 setae...............................24
24a(23b).
Head, pronotum, and mesepisternum with prominent surface sculpturing: northwestern United States.............................Rossianidae
24b.
Head, pronotum, and mesepisternum without prominent surface sculpturing ..................................................................................25
25a(24b).
Mandibles with uniform scraper blades or if toothed, SA-1 of metanotum with 25 setae on membrane between sclerites; case
cornucopia-shaped and mostly of sand grains; larvae 12 mm long .................................................................................Apataniidae
25b.
Mandibles not modified into scraper blades; case of vegetation or mineral materials, larvae often 12 mm long and with a transverse depression of the pronotum in most species...........................................................................................................Limnephilidae
F. Taxonomic Key to Families of Freshwater Megaloptera Larvae
1a.
Abdominal segments 1 – 7 with lateral filaments, last segment with a long, median filament (Fig. 85) ......................................Sialidae
1b.
Abdominal segments 1 – 8 with lateral filaments, last segment without a median filament, but with a pair of prolegs, each with a
pair of claws (Fig. 86) .......................................................................................................................................................Corydalidae
FIGURE 85 Sialidae; Sialis (dorsal view). FIGURE 86
Niqronia (dorsal view).
Corydalidae;
712
W. L. Hilsenhoff
G. Taxonomic Key to Adults of Aquatic, Semiaquatic, and Riparian Heteroptera
1a.
Antennae shorter than head, inserted beneath eyes and (except Ochteridae) not visible from above (Figs. 87, 88, 93, and 95) ............
......................................................................................................................................................................suborder Nepomorpha 2
1b.
Antennae longer than head, inserted in front of eyes and visible from above (Figs. 96 and 99 – 101)..................................................9
2a(1a).
Nepomorpha: Rostrum broad, blunt, and triangular, not distinctly segmented (Fig. 88); each front tarsus a 1-segmented scoop
fringed with setae (Fig. 89) ...................................................................................................................................................Corixidae
2b.
Rostrum cylindrical or cone-shaped, distinctly 3- or 4-segmented (Fig. 90); front tarsi not scooplike.................................................3
3a(2b).
Apex of abdomen with a long, slender, tubular respiratory appendage (Fig. 91) .....................................................................Nepidae
3b.
Apical respiratory appendages absent, or if present, short and flat (Fig. 92) .......................................................................................4
4a(3b).
Meso- and metathoracic legs with fringes of swimming hairs; ocelli absent; aquatic ..........................................................................5
4b.
Meso- and metathoracic legs without fringes or swimming hairs; ocelli usually present; riparian.......................................................8
5a(4a).
Dorsoventrally flattened, ovate insects; profemurs broad, raptorial (Fig. 93) .....................................................................................6
5b.
Elongate or hemispherical insects, not flattened dorsoventrally (Figs. 94, and 95); profemurs slender, similar to other legs. ..............7
6a(5a).
Length 18 mm; short, flat, straplike apical respiratory appendages present (Fig. 92); eyes protrude from margin of head.................
.....................................................................................................................................................................................Belostomatidae
6b.
Length 16 mm; apical respiratory appendages absent; eyes do not protrude from margin of head (Fig. 93) ....................Naucoridae
7a(5b).
Hemispherical (Fig. 94); length 3 mm ....................................................................................................................................Pleidae
7b.
Elongate (Fig. 95); length 5 mm ...................................................................................................................................Notonectidae
8a(4b).
Profemurs broad, raptorial; antennae concealed from above.........................................................................................Gelastocoridae
8b.
Profemurs slender, similar to other legs; antennae visible from above..................................................................................Ochteridae
9a(1b).
Membrane of hemelytra with 4 or 5 equal-sized cells (Fig. 96); metacoxae large, transverse; riparian .....Leptopodomorpha, Saldidae
9b.
Membrane of hemelytra without veins or with dissimilar sized cells; metacoxae small, conical; semiaquatic or riparian......................
...................................................................................................................................................................suborder Gerromorpha 10
10a(9b).
Gerromorpha: Claws of at least protarsi inserted before apex (Fig. 97)............................................................................................11
10b.
Claws of all tarsi inserted at apex (Fig. 98) ......................................................................................................................................12
11a(10a).
Metafemurs very long, greatly surpassing apex of abdomen ...................................................................................................Gerridae
11b.
Metafemurs short, not, or only slightly surpassing apex of abdomen.......................................................................................Veliidae
12a(10b).
Head as long as entire thorax, very slender, with eyes set about halfway to base (Fig. 99)............................................Hydrometridae
12b.
Head short and stout, eyes near posterior margin ............................................................................................................................13
13a(12b).
Head grooved ventrally to receive rostrum; tarsi 2-segmented; 2.5 mm long.......................................................................Hebridae
13b.
Head not grooved ventrally; tarsi 3-segmented; 2.5 mm long ........................................................................................................14
14a(13b).
Inner margins of eyes converge anteriorly (Fig. 100); femurs with 1 or more dorsal black spines distally ........................Mesoveliidae
14b.
Inner margins of eyes rounded (Fig. 101); femurs without black spines; riparian; western United States........................Macroveliidae
H. Taxonomic Key to Families of Freshwater Coleoptera
1. Key to Adult Water Beetles
1a.
Head formed into a snout or beak anteriorly; antennae geniculate (Fig. 104).................................................................Curculionidae
1b.
Head not formed into a beak or snout; antennae not geniculate.........................................................................................................2
2a(1b).
Two pairs of eyes, a dorsal and a ventral pair divided by sides of head; meso- and metathoracic legs short, extremely flat, with tarsi
folding fanlike.......................................................................................................................................................................Gyrinidae
2b.
One pair of eyes; meso- and metathoracic legs not extremely flat, with tarsi not folded fanlike..........................................................3
3a(2b).
Metacoxae expanded into large plates that cover 2 or 3 abdominal sterna and base of metafemurs (Fig. 105) ....................Haliplidae
17. Diversity and Classification of Insects and Collembola
FIGURE 87 Corixidae; Siqara (dorsal view), with right wings extended laterally. FIGURE 88 Corixidae;
head and prothorax of Siqara (ventral view) showing rostrum (R) and antenna (A). FIGURE 89 Corixidae;
pala of male Siqara. FIGURE 90 Notonectidae; head of Notonecta (ventral view) showing rostrum (R) and
antenna (A). FIGURE 91 Nepidae; apex of abdomen of Nepa (dorsal view). FIGURE 92 Belostomatidae;
apex of abdomen of Belostoma (dorsal view). FIGURE 93 Naucoridae; Pelocoris (dorsal view). FIGURE
94 Pleidae; Neoplea (lateral view). FIGURE 95 Notonectidae; Notonecta (dorsal view). FIGURE 96
Saldidae; Salda (dorsal view) showing membrane (M). FIGURE 97 Gerridae: protarsus of Gerris. FIGURE
98 Mesoveliidae; protarsus of Mesovelia. FIGURE 99 Hydrometridae; Hydrometra (dorsal view).
FIGURE 100 Mesoveliidae; head of Mesovelia (dorsal view). FIGURE 101 Macroveliidae; head of
Macrovelia (dorsal view).
713
714
W. L. Hilsenhoff
FIGURE 102 Neuroptera Larva (Sisyridae); Climacia (dorsal view). FIGURE 103
(Pyralidae); Nymphula (lateral view).
Lepidoptera Larva
3b.
Metacoxae not expanded into large plates .........................................................................................................................................4
4a(3b).
Prosternum with a postcoxal process that extends posteriorly to mesocoxae (Fig. 106); first visible abdominal sternum completely
divided by metacoxal process (Fig. 106) .............................................................................................................................................5
4b.
Prosternum with postcoxal process absent or short; first visible abdominal sternum extending for its entire breadth behind metacoxae (Fig. 107) .................................................................................................................................................................................8
5a(4a).
Base of pronotum much narrower than base of elytra (Fig. 108); metatarsi rounded and not fringed with long setae; large 10 mm
long; western mountains of the United States...................................................................................................................Amphizoidae
5b.
Base of pronotum subequal in width to base of elytra; metatarsi flattened and fringed with long setae or beetles 2 mm long..........6
6a(5b).
Anterior of prosternum, its postcoxal process and metasternum in same plane (Fig. 109); pro- and mesotarsi distinctly 5-segmented,
segment 4 as long as 3 ........................................................................................................................................................................7
6b.
Anterior of prosternum greatly depressed and not in same plane as its postcoxal process and metasternum (Fig. 110); pro- and
mesotarsi appear to be 4-segmented .......................................................................................................................Dytiscidae (in part)
7a(6a).
Prosternal process pointed or nearly so (Figs. 106); no curved spur or hooked apex on protibiae; 4 mm long ....Dytiscidae (in part)
7b.
Prosternal process truncate or rounded apically (Fig. 111); protibiae with curved spur or hooked apex (Fig. 112) or beetles 3 mm
long; eastern, central United States .......................................................................................................................................Noteridae
8a(4b).
Antennae short, club-shaped, with segment 4 or 6 modified to form a cupule (Fig. 113); maxillary palps usually longer than antennae .....................................................................................................................................................................................................9
8b.
Antennae pectinate or filiform (Figs. 116, 117), usually longer than maxillary palps .......................................................................12
9a(8a).
Antennae with 5 segments past cupule; less than 2.5 mm long .........................................................................................Hydraenidae
9b.
Antennae with 3 segments past cupule (Fig. 113); 1.5 – 40.0 mm long..............................................................................................10
10a(9b).
Pronotum with 7 longitudinal grooves, including marginal grooves (Fig. 114); 2.0 – 7.1 mm long ..................................Helophoridae
10b.
Pronotum without 7 longitudinal grooves ........................................................................................................................................11
11a(10b).
Pronotum much narrower than elytral base and scutellum very small (Fig. 115); pronotum granular or with large punctures;
2.2 – 5.3 mm long .............................................................................................................................................................Hydrochidae
Pronotum not much narrower than base of elytra, or scutellum elongate; pronotum not granulate, large punctures lacking;
1.5 – 42.0 mm long.........................................................................................................................................................Hydrophilidae
17. Diversity and Classification of Insects and Collembola
715
FIGURE 104 Curculionidae; head (lateral view). FIGURE 105 Haliplidae; metathorax and
abdomen of Haliplus (ventral view) showing metacoxal plate (CP). FIGURE 106 Dytiscidae;
Aqabus (ventral view) showing prosternal process (PP), first abdominal sternum (A-1), and metacoxal
process (MP). FIGURE 107 Hydrophilidae; Tropisternus (ventral view) showing first abdominal
sternum (A-1). FIGURE 108 Amphizoidae; Amphizoa (dorsal view). FIGURE 109 Dytiscidae;
Aqabus (lateral view) showing prosternum (PS), prosternal process (PP), metasternum (MS), procoxa
(PC), and mesocoxa (MC). FIGURE 110 Dytiscidae; Hydroporus (lateral view) showing
prosternum (PS), prosternal process (PP), metasternum (MS), procoxa (PC), and mesocoxa (MC).
12a(8b).
Antennae short with pectinate club (Fig. 116); 5.0 mm long ............................................................................................Dryopidae
12b.
Antennae slender, filiform; (Fig. 117); 4.5 mm long ......................................................................................................................13
13a(12b).
Extremely small, 1.5 mm long (Fig. 118); all tarsi 3-segmented; southwest United States, north to Idaho ...............Hydroscaphidae
13b.
Larger, 1.7 mm long; all tarsi 5-segmented ...........................................................................................................................Elmidae
716
W. L. Hilsenhoff
FIGURE 111 Noteridae; thorax of Hydrocanthus (ventral view) showing prosternal process (PP) and
metasternal plate (MP). FIGURE 112 Noteridae; prothoracic leg of Hydrocanthus showing profemur (F), tibia
(TI) and tarsus (TA). FIGURE 113 Hydrophilidae; antenna of Tropisternus showing cupule (C). FIGURE 114
Helophoridae; pronotum of Helophorus showing longitudinal grooves. FIGURE 115 Hydrochidae;
Hydrochus (dorsal view). FIGURE 116 Dryopidae; right antenna and eye of Helichus (dorsal view). FIGURE
117 Elmidae; Stenelmis (dorsal view). FIGURE 118 Hydroscaphidae; Hydroscapha (dorsal view).
2. Key to Larval Water Beetles
1a.
Tarsi with 2 claws ..............................................................................................................................................................................2
1b.
Tarsi with 1 claw................................................................................................................................................................................5
2a(1a).
Abdomen with 4 conspicuous hooks on last segment; abdominal segments with at least 8 pairs of lateral filaments (Fig. 119)
Gyrinidae
2b.
No hooks on last abdominal segment; if lateral abdominal filaments are present, there are only 6 pairs ............................................3
3a(2b).
Abdominal and thoracic terga flattened and expanded laterally (Fig. 120); western mountains of the United States........Amphizoidae
3b.
Abdominal and thoracic terga not flattened and expanded laterally ...................................................................................................4
4a(3b).
Urogomphi shorter than last abdominal segment; legs short, stout, adapted for digging (Fig. 121); mandibles short, adapted for
chewing; eastern, central United States .................................................................................................................................Noteridae
4b.
Urogomphi usually longer than last abdominal segment (Fig. 122); if shorter, legs are elongate with setal fringe for swimming;
mandibles elongate, pointed, for piercing .............................................................................................................................Dytiscidae
5a(1b).
Legs distinctly 5-segmented; abdomen terminating in 1 or 2 long filaments (Figs. 123, and 124) .........................................Haliplidae
5b.
Legs apparently 4-segmented; abdomen not terminating in long filaments .........................................................................................6
6a(5b).
Mandibles large, readily visible from above (Fig. 125)...................................................................................................Hydrophilidae
6b.
Mandibles not readily visible from above ...........................................................................................................................................7
7a(6b).
Antennae long, filiform, as long as head and thorax combined (Fig. 126) ...............................................................................Scirtidae
7b.
Antennae much shorter than head and thorax combined ...................................................................................................................8
8a(7b).
Body oval and extremely flat; head completely concealed from dorsal view (Figs. 127, and 128) ......................................Psephenidae
8b.
Body elongate and round or triangular in cross section; head exposed, except mostly concealed in Lampyridae ................................9
9a(8b).
Each thoracic and abdominal segment covered by a flat, platelike sclerite dorsally; prothoracic plate mostly or completely concealing head from above (Fig. 129) ..........................................................................................................................................Lampyridae
17. Diversity and Classification of Insects and Collembola
717
FIGURE 119 Gyrinidae; Dineutus (dorsal view). FIGURE 120 Amphizoidae; Amphizoa (dorsal view).
FIGURE 121 Noteridae; Hydrocanthus (dorsal view). FIGURE 122 Dytiscidae; Aqabus (dorsal view)
showing urogomphi (U). FIGURE 123 Haliplidae; Haliplus (dorsal view). FIGURE 124 Haliplidae;
Peltodytes (lateral view). FIGURE 125 Hydrophilidae; Tropisternus (dorsal view). FIGURE 126
Scirtidae; Cyphon (dorsal view). FIGURE 127 Psephenidae; Ectopria (dorsal view). FIGURE 128
Psephenidae; Psephenus (dorsal view). FIGURE 129 Lampyridae (lateral view). FIGURE 130
Chrysomelidae; Donacia (lateral view). FIGURE 131 Ptilodactylidae; last abdominal segment of
Anchytarsus (lateral view). FIGURE 132 Elmidae; Stenelmis (lateral view). FIGURE 133 Elmidae; last
abdominal tergum of Stenelmis. FIGURE 134 Lutrochidae; last abdominal tergum of Lutrochus.
9b.(8a).
Thoracic and abdominal segments not covered by flat, platelike sclerites; head visible from above ..................................................10
10a(9b).
All terga rounded and pale; grublike larvae with 2 spines on last abdominal tergum (Fig. 130) ....................................Chrysomelidae
10b.
Body elongate and sclerotized; no spines on last abdominal tergum (Figs. 131, and 132).................................................................11
11a(10b).
Abdominal sterna with distinct tufts of gills, either on sterna 1 – 7 or on last segment and lacking and operculum (Fig. 131); eastern,
United States and California..........................................................................................................................................Ptilodactylidae
718
W. L. Hilsenhoff
11b.
Abdominal gills, if present, on last abdominal segment and covered by a ventral operculum (Fig. 132) ...........................................12
12a(11b).
Last abdominal tergum bifid or notched apically (Fig. 133) .....................................................................................................Elmidae
12b.
Last abdominal tergum rounded apically (Fig. 134); East, Southwest. ...............................................................................Lutrochidae
I. Taxonomic Key to Families of Freshwater Diptera Larvae
1a.
Larvae apparently 7-segmented; first 6 segments each with a prominent ventral sucker (Fig. 135) ..........Nematocera, Blephariceridae
1b.
Larvae with more than 7 apparent segments; without 6 ventral suckers .............................................................................................2
2a(1b).
Head capsule completely sclerotized and fully visible (Figs. 147 – 159); mandibles opposed, moving in a horizontal plane, and usually with two or more apical teeth; body never flattened and with a posterior spiracular chamber margined with long, soft hairs.
.....................................................................................................................................................................................Nematocera 12
2b.
Sclerotized head capsule absent (Figs 138 – 140, 142, 143, 145, and 146), or incomplete behind (Figs. 136 and 137), and retracted
at least partially into thorax; mandibles parallel, without secondary apical teeth, and moving in a vertical plane or in a partially
sclerotized retracted head may be opposed and moving in a horizontal plane ....................................................................................3
3a(2b).
Mandibles opposed, moving in a horizontal plane, and with two or more apical teeth; larvae truncate anteriorly with head capsule
retracted into thorax and incomplete posteriorly (Figs. 136 and 137)...............................................................Nematocera, Tipulidae
3b.
Mandibles parallel, moving vertically, and without secondary apical teeth; larvae narrowed anteriorly with head capsule lacking or
poorly developed and incompletely sclerotized or body terminating in a long respiratory tube (Fig. 139) or a posterior spiracular
chamber margined with long, soft hairs ...........................................................................................................................Brachycera 4
4a(3b).
Brachycera: Head mostly visible, truncate in shape (Fig. 138); body somewhat flattened; posterior spiracular chamber margined
with long, soft hairs; integument covered with shiny calcium carbonate crystals ............................................................Stratiomyidae
4b.
Head mostly retracted into thorax and elongate, or indistinguishable; body nearly circular in cross section; without a posterior
spiracular chamber margined with long, soft hairs; integument lacking calcium carbonate crystals....................................................5
5a(4b).
Larvae with a partially retractile caudal respiratory tube at least one-half as long as body (Fig. 139) ...................................Syrphidae
5b.
Larvae without a long respiratory tube, if a short tube is present, it is divided apically ......................................................................6
6a(5b).
Body terminating in a short tube that is divided apically (Fig. 140), or in a pair of spines ..................................................Ephydridae
6b.
Body not terminating in a short tube or a pair of spines .....................................................................................................................7
7a(6b).
Caudal spiracular disk with palmate hairs and surrounded by 8 – 10 lobes, some of which may be very short (Fig. 141); body wrinkled ...................................................................................................................................................................................Sciomyzidae
7b.
Caudal spiracular disk without palmate hairs, if surrounded by lobes, body not wrinkled .................................................................8
8a(7b).
Abdomen without distinct prolegs, paired ventral pseudopods, or terminal processes except a single spine, but with a girdle of 6 or
more pseudopods on each segment (Fig. 142) ......................................................................................................................Tabanidae
8b.
Abdomen with distinct prolegs, paired pseudopods, or paired terminal processes present ..................................................................9
9a(8b).
Body terminating in a spiracular pit surrounded by 4 pointed lobes (Fig. 143) ............................................................Dolichopodidae
9b.
Body not terminating in a spiracular pit with 4 lobes .......................................................................................................................10
10a(9b).
Body terminating in a pair of ciliated, divergent processes (Fig. 144); abdomen with 8 pairs of ventral prolegs .................Athericidae
10b.
Terminal processes, if present, not ciliated; abdomen with or without prolegs .................................................................................11
11a(10b).
Some external head structure visible, with palps and antennae usually present; abdomen usually with ventral prolegs and elongate,
paired, terminal appendages (Fig. 145) or a bulbous segment ..............................................................................................Empididae
11b.
No visible external head structure; abdomen often with ventral pseudopods and usually short, paired, terminal appendages (Fig.
146) ......................................................................................................................................................................................Muscidae
12a(2a).
Nematocera: Prolegs absent (Figs. 147, 150, and 151) .....................................................................................................................13
12b.
Prolegs present at one or both ends of body or on abdominal segments (Figs. 152 – 159).................................................................17
13a(12a).
Thoracic segments fused and distinctly thicker than abdomen (Fig. 147) .........................................................................................14
13b.
Thoracic segments not fused and about equal to abdomen in diameter (Figs. 150 and 151).............................................................16
14a(13a).
Antennae prehensile, with long, strong apical spines (Fig. 148) ........................................................................................................15
17. Diversity and Classification of Insects and Collembola
719
FIGURE 135 Blephariceridae; Blepharicera (ventral view). FIGURE 136 Tipulidae; head of Limonia
(dorsal view). FIGURE 137 Tipulidae; head of Hexatoma (dorsal view). FIGURE 138 Stratiomyidae; head
of Odontomyia (dorsal view). FIGURE 139 Syrphidae; Eristalis (lateral view). FIGURE 140 Ephydridae
(lateral view). FIGURE 141 Sciomyzidae; spiracular disc of Sepedon. FIGURE 142 Tabanidae; Chrysops
(lateral view). FIGURE 143 Dolicopodidae (lateral view). FIGURE 144 Athericidae; terminal segments of
Atherix (dorsal view). FIGURE 145 Empididae (lateral view). FIGURE 146 Muscidae; Limnophora
(lateral view). FIGURE 147 Culicidae; Anopheles (dorsal view). FIGURE 148 Chaoboridae; head of
Chaoborus (lateral view) showing antenna (A). FIGURE 149 Culicidae; head of Coquillettidia (dorsal view)
showing antenna (A). FIGURE 150 Psychodidae; Psychoda (dorsal view). FIGURE 151 Ceratopogonidae;
Palpomyia (dorsal view). FIGURE 152 Nymphomyiidae; Palaeodipteron (lateral view).
14b.
Antennae not prehensile, lacking long apical spines (Fig. 149)...............................................................................................Culicidae
15a(14a).
Antennae close together at base; a row of spinose setae on dorsolateral margin of head ................................................Corethrellidae
15b.
Antennae at anterolateral margins of head; head without spinose setae dorsolaterally .....................................................Chaoboridae
720
W. L. Hilsenhoff
FIGURE 153 Deuterophlebiidae; Deuterophlebia (dorsal view). FIGURE 154 Dixidae; Dixella (lateral
view). FIGURE 155 Ptychopteridae; Ptychoptera (lateral view). FIGURE 156 Simuliidae; Simulium (lateral
view). FIGURE 157 Tanyderidae; Protoplasa (lateral view). FIGURE 158 Chironomidae; Chironomus
(lateral view). FIGURE 159 Thaumaleidae; Thaumalea (lateral view).
16a(13b).
Thoracic and abdominal segments each distinctly divided into 2 or 3 annuli, with sclerotized dorsal plates on some annuli (Fig. 150)
Psychodidae
16b.
No secondary annulations on abdominal segments (Fig. 151) .....................................................................Ceratopogonidae (in part)
17a(12b).
Prolegs on intermediate body segments (Figs. 152 – 155) ..................................................................................................................18
17b.
Prolegs on anterior and / or posterior ends of body only (Figs. 156 – 159) .........................................................................................21
18a(17a).
Seven or eight pairs of distinct prolegs on abdomen (Figs. 152 and 153)..........................................................................................19
18b.
Two or three pairs of weak prolegs on abdomen (Figs. 154 and 155)...............................................................................................20
19a(18a).
Eight pairs of slender, ventrally projecting prolegs (Fig. 152); PQ, NB, ME ...............................................................Nymphomyiidae
19b.
Seven pairs of stout, ventrolaterally projecting prolegs (Fig. 153); western mountains.............................................Deuterophlebiidae
20a(18b).
Paired ventral prolegs on abdominal segment 1 and usually also on segment 2; posterior end of body with 2 pairs of fringed
processes (Fig. 154) ..................................................................................................................................................................Dixidae
20b.
Paired ventral prolegs on abdominal segments 1, 2, and 3; body terminating in a long respiratory tube (Fig. 155) .......Ptychopteridae
21a(17b).
A single proleg present only on prothorax; posterior of abdomen swollen and terminating in a ring of numerous small hooks (Fig.
156) .....................................................................................................................................................................................Simuliidae
21b.
Posterior prolegs usually present; posterior of abdomen not swollen and terminating in a ring of hooks .........................................22
22a(21b).
Only posterior prolegs present (Fig. 157) .........................................................................................................................................23
22b.
Anterior and usually posterior prolegs present (Figs. 158 and 159) ..................................................................................................24
23a(22a).
Long filamentous processes arising from last two abdominal segments and from prolegs (Fig. 157); Eastern and Western United
States.................................................................................................................................................................................Tanyderidae
23b.
Last two abdominal segments without long filamentous processes...............................................................Ceratopogonidae (in part)
24a(22b).
Body covered with long, strong spines .........................................................................................................Ceratopogonidae (in part)
24b.
Body at most covered with setae ......................................................................................................................................................25
25a(24b).
At least one pair of prolegs separated distally (Fig. 158); stalked spiracles lacking.........................................................Chironomidae
25b.
Prolegs unpaired (Fig. 159); short, stalked spiracles dorsolaterally on the prothorax; mountain streams........................Thaumaleidae
J. Taxonomic Key to Families of Semiaquatic Collembola
1a.
Body somewhat globular; thorax and abdomen indistinctly segmented (Fig. 160) ...........................................................Sminthuridae
1b.
Body elongate; thorax and abdomen distinctly segmented (Fig. 161) .................................................................................................2
17. Diversity and Classification of Insects and Collembola
721
FIGURE 160 Sminthuridae (lateral view) showing furcula (F) and collophore (C). FIGURE 161 Poduridae;
Podura (lateral view) showing furcula (F) and collophore (C). FIGURE 162 Poduridae; furcula of Podura
(dorsal view) showing manubrium (MA), dens (D), and mucro (MU). FIGURE 163 Isotomidae anterior
portion (lateral view). [Figures 158 – 161 redrawn from Peckarsky et al. (1990).]
2a(1b).
Mouthparts directed downward (Fig. 161); furcula distinctly convergent apically (Fig. 162); first abdominal segment with dorsal
setae .....................................................................................................................................................................................Poduridae
2b.
Mouthparts directed forward (Fig. 163); furcula not distinctly convergent apically; first abdominal segment lacking dorsal setae .......
...........................................................................................................................................................................................Isotomidae
LITERATURE CITED
Adler, P. H., Kim, K. C. 1986. The blackflies of Pennsylvania (Simuliidae, Diptera). Bionomics, taxonomy, and distribution. Pennsylvania State University Agricultural Experiment Station Bulletin
856, 88 pp.
Agriculture Canada, Research Branch. 1981. Manual of Nearctic
Diptera Vol. 1. Monograph No. 27. Canadian Government
Publishing Centre, Supply and Services Canada, Hull, Que.,
Canada, vi 674 pp.
Agriculture Canada, Research Branch. 1987. Manual of Nearctic
Diptera Vol. 2. Monograph No. 28. Canadian Government Publishing Centre, Supply and Services Canada, Hull, Que., Canada,
vi 658 pp.
Alarie, Y. 1991. Description of larvae of 17 Nearctic species of Hydroporus Clairville (Coleoptera: Dytiscidae: Hydroporinae)
with an analysis of their phylogenetic relationships. Canadian
Entomologist 123:627 – 704.
Alarie, Y. 1992. Descriptions of the larval stages of Hydroporus
(Heterosternuta) Cocheconis Fall 1917 (Coleoptera: Dytiscidae:
Hydroporinae) with a key to the known larvae of Heterosternuta Strand. Canadian Entomologist 124:827 – 840.
Alarie, Y. 1993. A systematic review of the North American species
of the Oreodytes alaskanus clade (Coleoptera: Dytiscidae: Hydroporinae). Canadian Entomologist 125:847 – 867.
Alarie, Y. 1997. Taxonomic revision and phylogenetic analysis of the
genus Oreodytes Seidlitz (Coleoptera: Dytiscidae: Hydroporinae) based on larval morphology. Canadian Entomologist 129:
399 – 503.
Alarie, Y., Harper, P. P., Maire, M. 1989. Rearing dytiscid beetles
(Coleoptera: Dytiscidae). Entomologica Basiliensia 13:147 – 149.
Alarie, Y., Harper, P. P., Roughley, R. E. 1990. Description of the larvae of eleven Nearctic species of Hygrotus Stephens (Coleoptera:
Dytiscidae: Hydroporinae) with an analysis of their phyletic relationships. Canadian Entomologist 122:985 – 1035.
Allen, R. K. 1973. Generic revisions of mayfly nymphs. 1. Traverella
in North and Central America (Leptophlebiidae). Annals of the
Entomological Society of America 66:1287 – 1295.
Allen, R. K. 1978. The nymphs of North and Central American
Leptohyphes (Ephemeroptera: Tricorythidae). Annals of the Entomological Society of America 71:537 – 558.
Allen, R. K., Brusca, R. C. 1978. Generic revisions of mayfly nymphs
II. Thraulodes in North and Central America (Leptophlebiidae).
Canadian Entomologist 110:413 – 433.
Allen, R. K., Edmunds, G. F., Jr. 1959. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) I. The subgenus
Timpanoga. Canadian Entomologist 91:51 – 58.
Allen, R. K., Edmunds, G. F., Jr. 1961a. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) II. The subgenus
Caudatella. Annals of the Entomological Society of America
54:603 – 612.
Allen, R. K., Edmunds, G. F., Jr. 1961b. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) III. The subgenus Attenuatella. Journal of the Kansas Entomological Society 34:161 – 173.
Allen, R. K., Edmunds, G. F., Jr. 1962a. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) IV. The subgenus Dannella. Journal of the Kansas Entomological Society
35:333 – 338.
Allen, R. K., Edmunds, G. F., Jr. 1962b. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) V. The subgenus
Drunella in North America. Miscellaneous Publications of the
Entomological Society of America 3:147 – 187.
Allen, R. K., Edmunds, G. F., Jr. 1963a. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) VI. The subgenus Serratella in North America. Annals of the Entomological
Society of America 56:583 – 600.
Allen, R. K., Edmunds, G. F., Jr. 1963b. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) VII. The subgenus Eurylophella. Canadian Entomologist 95:597 – 623.
Allen, R. K., Edmunds, G. F., Jr. 1965. A revision of the genus
Ephemerella (Ephemeroptera: Ephemerellidae) VIII. The subgenus Ephemerella in North America. Miscellaneous Publications of the Entomological Society of America 4:244 – 282.
Allen, R. K., Edmunds, G. F., Jr. 1976. A revision of the genus
Ametropus in North America (Ephemeroptera: Ametropodidae). Journal of the Kansas Entomological Society 49:625 – 635.
Allen, R. K., Murvosh, C. M. 1987. Mayflies (Ephemeroptera:
Tricorythidae) of the southwestern United States and northern
Mexico. Annals of the Entomological Society of America
80:35 – 40.
Andersen, N. M. 1990. Phylogeny and taxonomy of water striders,
genus Aquarius Schellenberg (Insecta, Hemiptera, Gerridae),
with a new species from Australia. Steenstrupia 16:37 – 81.
Andersen, N. M. 1993. Classification, phylogeny, and zoogeography
of the pond skater genus Gerris Fabricius (Hemiptera: Gerridae). Canadian Journal of Zoology 71:2473 – 2508.
Andersen, N. M., Spence, J. R. 1992. Classification and phylogeny of
722
W. L. Hilsenhoff
the Holarctic water strider genus Limnoporus Stal (Hemiptera:
Gerridae). Canadian Journal of Zoology 70:753 – 785.
Anderson, L. D. 1932. A monograph of the genus Metrobates. University of Kansas Science Bulletin 20:297 – 311.
Anderson, N. H., Sedell, J. R. 1979. Detritus processing by macroinvertebrates in stream ecosystems. Annual Review of Entomology 24:351 – 377.
Anderson, R. D. 1962. The Dytiscidae (Coleoptera) of Utah: keys,
original citation, types, and Utah distribution. Great Basin Naturalist 22: 54 – 75.
Anderson, R. D. 1970. A revision of the Nearctic representatives of
Hygrotus (Coleoptera: Dytiscidae). Annals of the Entomological
Society of America 64:503 – 512.
Anderson, R. D. 1975. A revision of the Nearctic species of Hygrotus
groups II and III (Coleoptera: Dytiscidae). Annals of the Entomological Society of America 69:577 – 584.
Anderson, R. D. 1983. Revision of the Nearctic species of Hygrotus
groups IV, V, and VI (Coleoptera: Dytiscidae). Annals of the Entomological Society of America 76:173 – 196.
Archangelsky, M. 1998. Phylogeny of Hydrophiloidea (Coleoptera:
Staphyliniformia) using characters from adult and preimaginal
stages. Systematic Entomology 23:9 – 24.
Bae, Y. J., McCafferty, W. P. 1991. Phylogenetic systematics of the
Potamanthidae (Ephemeroptera). Transactions of the American
Entomological Society 117(3,4):1 – 143.
Bae, Y. J., McCafferty, W. P. 1998. Phylogenetic systematics and biogeography of the Neoephemeridae (Ephemeroptera: Pannota).
Aquatic Insects 20:35 – 68.
Barr, C. B., Chapin, J. B. 1988. The aquatic Dryopoidea of Louisiana
(Coleoptera: Psephenidae, Dryopidae, Elmidae). Tulane Studies
in Zoology and Botany 26:89 – 164.
Baumann, R. W., Gaufin, A. R., Surdick, R. F. 1977. The stoneflies
(Plecoptera) of the Rocky Mountains. Memoirs of the American
Entomological Society 31, ii 208 pp.
Baumann, R. W., Stark, B. P. 1980. Hesperoperla hoguei, a new
species of stonefly from California (Plecoptera: Perlidae). Great
Basin Naturalist 40:63 – 67.
Bay, E. C. 1974. Predator – prey relationships among aquatic insects.
Annual Review of Entomology 19:441 – 453.
Bednarik, A. F., Edmunds, G. F. Jr. 1980. Descriptions of larval Heptagenia from the Rocky Mountain region. (Ephemeroptera:
Heptageniidae). Pan-Pacific Entomologist 56:51 – 62.
Bednarik, A. F., McCafferty, W. P. 1979. Biosystematic revision of the
genus Stenonema (Ephemeroptera: Heptageniidae). Canadian
Bulletin of Fisheries and Aquatic Sciences 201, vi 73 pp.
Bennefield, B. L. 1965. A taxonomic study of the subgenus Ladona
(Odonata: Libellulidae). University of Kansas Science Bulletin
45:361 – 389.
Bennett, D. V., Cook, E. F. 1981. The semiaquatic Hemiptera of Minnesota (Hemiptera: Heteroptera). Technical Bulletin 332 Agricultural Experiment Station, University of Minnesota, 59 pp.
Berg, C. O., Knudson, L. 1978. Biology and systematics of the
Sciomyzidae. Annual Review of Entomology 23:239 – 258.
Berge Henegouwen, van, A. 1986. Revision of the European species
of Anacaena Thomson (Coleoptera: Hydrophilidae). Entomologica Scandinavica 17:393 – 407.
Bergman, E. A., Hilsenhoff, W. L. 1978. Baetis (Ephemeroptera:
Baetidae) of Wisconsin. Great Lakes Entomologist 11:125 – 135.
Berner, L. 1956. The genus Neoephemera in North America
(Ephemeroptera: Neoephemeridae). Annals of the Entomological Society of America 49:33 – 42.
Berner, L. 1959. A tabular summary of the biology of North American mayfly nymphs (Ephemeroptera). Bulletin of the Florida
State Museum 4:1 – 58.
Berner, L., Pescador, M. L. 1988. The mayflies of Florida. Revised
edition., Uni. Press of Florida, Gainesville, FL, 415 pp.
Bilyj, B. 1988. A taxonomic review of Guttipelopia (Diptera: Chironomidae). Entomologica Scandinavica 19:1 – 26.
Bistrom, O. 1996. Taxonomic revision of the genus Hydrovatus
Motschulsky (Coleoptera, Dytiscidae). Entomologica Basiliensia
19:57 – 584.
Bobb, M. L. 1974. The insects of Virginia: No. 7. The aquatic and
semi-aquatic Hemiptera of Virginia. Research Division Bulletin
87, Virginia Polytechnic Institute and State University, Blacksburg, iv 196 pp.
Bode, R. W. 1983. Larvae of North American Eukiefferiella and
Tvetenia (Diptera: Chironomidae). Bulletin 452, New York
State Museum, v 40 pp.
Boesel, M. W. 1985. A brief review of the genus Polypedilum in
Ohio, USA with keys to known stages of species occurring in
northeastern USA. Ohio Journal of Science 85:245 – 262.
Borkent, A. 1979. Systematics and bionomics of the species of the subgenus Schadonophasma Dyar and Shannon (Chaoborus, Chaoboridae, Diptera). Quaestiones Entomologicae 15:122 – 255.
Borkent, A. 1984. The systematics and phylogeny of the Stenochironomus complex (Xestochironomus, Harrisius, and Stenochironomus) (Diptera: Chironomidae). Memoirs of the Entomological Society of Canada 128, 269 pp.
Bowles, D. E. 1995. A new species of Austrotinodes (Trichoptera: Ecnomidae) from Texas. Journal of the New York Entomological
Society 103:155 – 161.
Bratt, A. D., Knutson, L. V., Foote, B. A., Berg, C. O. 1969. Biology
of Pherbellia (Diptera: Sciomyzidae). Memoirs of the Cornell
University Agricultural Experiment Station 404, 247 pp.
Breeland, S. G., Loyless, T. M. 1982. Illustrated keys to the mosquitoes of Florida. Adult females and fourth stage larvae. Journal
of the Florida Anti-mosquito Association 53:63 – 84.
Brigham, A. R., Brigham, W. U., Gnilka, A. Eds. 1982. The aquatic
insects and oligochaetes of North and South Carolina. Midwest
Aquatic Enterprises, Mahomet, IL, xi 837 pp.
Brigham, W. U. 1982a. Megaloptera. in: Brigham, A. R., Brigham,
W. U. Gnilka, A. Eds. The aquatic insects and oligochaetes of
North and South Carolina. Midwest Aquatic Enterprises, Mahomet, IL, pp. 7.1 – 7.12.
Brigham, W. U. 1982b. Aquatic Neuroptera. in: Brigham, A. R.,
Brigham, W. U., Gnilka, A. Eds. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 8.1 – 8.4.
Brigham, W. U. 1982c. Aquatic Coleoptera. in Brigham, A. R.,
Brigham, W. U., Gnilka, A. Eds. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 10.1 – 10.136.
Brittain, J. E. 1982. Biology of mayflies. Annual Review of Entomology 27:119 – 147.
Brooks, A. R., Kelton, L. A. 1967. Aquatic and semiaquatic Heteroptera of Alberta, Saskatchewan, and Manitoba (Hemiptera).
Memoirs of the Entomological Society of Canada 51. 92 pp.
Brown, H. P. 1972. Aquatic dryopoid beetles (Coleoptera) of the
United States. Biota of Freshwater Ecosystems Identification
Manual 6. U.S. Environmental Protection Agency Water Pollution Control Research Series 18050 ELDO4/72, ix 82 pp.
Brown, H. P. 1987. Biology of riffle beetles. Annual Review of Entomology 32:253 – 273.
Burger, J. F. 1977. The biosystematics of immature Arizona Tabanidae (Diptera). Transactions of the American Entomological Society 103:145 – 268.
Burian, S. K., Novak, M. A., Bode, R. W., Abele, L. 1997. New
record of Brachycercus maculatus Berner (Ephemeroptera:
17. Diversity and Classification of Insects and Collembola
Caenidae) from New York and a key to larvae of northeastern
species. Great Lakes Entomologist 30:85 – 88.
Burks, B. D. 1953. The mayflies or Ephemeroptera of Illinois. Bulletin of the Illinois Natural History Survey 26:1 – 216.
Calabrese, D. 1974. Keys to the adults and nymphs of the species of
Gerris Fabricius occurring in Connecticut. Memoirs, Connecticut Entomological Society 1974: 227 – 266.
Centerbury, L. E. 1978. Studies of the genus Sialis (Sialidae: Megaloptera) in eastern North America. Ph.D. Dissertation, University of Louisville, KY, x 93 pp.
Carle, F. L. 1980. A new Lanthus (Odonata: Gomphidae) from eastern North America with adult and nymphal keys to American
octogomphines. Annals of the Entomological Society of America 73: 172 – 179.
Carle, F. L. 1992. Ophiogomphus (Ophionurus) australis spec. nov.
from the Gulf coast of Louisiana, with larval and adult keys to
American Ophiogomphus (Anisoptera: Gomphidae). Odonatologica 21:141 – 152.
Carpenter, S. J., LaCasse, W. J. 1955. Mosquitoes of North America.
Uni. of California Press, Berkeley, CA, vi 360 pp.
Chapin, J. W. 1978. Systematics of Nearctic Micrasema (Trichoptera:
Brachycentridae). Ph.D. Dissertation, Clemson University,
Clemson, SC, xiii 136 pp.
Chapman, H. C. 1958. Notes on the identity, habitat and distribution of some semiaquatic Hemiptera of Florida. Florida Entomologist 41:117 – 124.
Cheary, B. S. 1971. The biology, ecology, and systematics of the
genus Laccobius (Laccobius) (Coleoptera: Hydrophilidae) of
the new world. Ph.D. Dissertation, University of California,
Riverside, xv 178 pp. (privately printed).
Check, G. R. 1982. A revision of the North American species of
Callibaetis (Ephemeroptera: Baetidae). Ph.D. Dissertation, University of Minnesota, Minneapolis, 164 pp.
Chen, Y. E. 1992. The larva and pupa of Apatania praevolans
Morse (Trichoptera: Limnephilidae), with a key to described
larvae of North American species of Apatania. Aquatic Insects
14:49 – 55.
Cheng, L., Fernando, C. H. 1970. The waterstriders of Ontario.
Miscellaneous Life Science Publications of the Royal Ontario
Museum, 23 pp.
Christiansen, K., Bellinger, P. 1980. The Collembola of North America north of the Rio Grande. A taxonomix analysis. Grinnell
College, IA, iii 1322 pp.
Clifford, H. F. 1991. Aquatic invertebrates of Alberta. Uni. of Alberta Press, Edmonton, 538 pp.
Cook, E. F. 1956. The Nearctic Chaoborinae (Diptera: Culicidae).
Technical Bulletin 218, University of Minnesota Agricultural
Experiment Station, 102 pp.
Corbet, P. S. 1962. A biology of dragonflies. H.F. & G. Witherby
Ltd., London, xvi 247 pp.
Corbet, P. S. 1980. Biology of Odonata. Annual Review of Entomology, 25:189 – 217.
Courtney, G. W. 1990. Revision of the Nearctic mountain midges
(Diptera: Deuterophlebiidae). Journal of Natural History
2:81 – 118.
Courtney, G. W. 1994. Biosystematics of the Nymphomyiidae
(Insecta: Diptera): life history, morphology and phylogenetic
relationships. Smithsonian Contributions to Zoology 550:1 – 41.
Currie, D. C. 1986. An annotated list of and keys to the immature
black flies of Alberta (Diptera: Simuliidae). Memoirs of the Entomological Society of Canada 134:1 – 90.
Cuyler, R. D. 1958. The larvae of Chauliodes Latreille (Megaloptera:
Corydalidae). Annals of the Entomological Society of America
51:582 – 586.
723
Daigle, J. J. 1991a. A new key to the larvae of North American Somatochlora. Argia 3(3):9 – 10.
Daigle, J. J. 1991b. Florida damselflies (Zygoptera): A species key to
the aquatic larval stages. Florida Department of Environmental
Regulation Technical Series 11:1 – 12.
Daigle, J. J. 1992. Florida dragonflies (Anisoptera): A species key to
the aquatic larval stages. Florida Department of Environmental
Regulation Technical Series 12:1 – 29.
Darsie, R. F., Ward, R. A. 1981. Identification and geographical distribution of the mosquitoes of North America, north of Mexico.
Mosquito Systematics Supplement 1, American Mosquito Control Association, 313 pp.
Davis, J. R. 1996. Naucoridae of Texas. Southwestern Naturalist
41:1 – 26.
Day, W. C. 1956. Ephemeroptera. in: Usinger, R. L., Ed. Aquatic insects of California with keys to North American genera and
California species. Uni. of California Press, Berkeley., CA,
pp. 79 – 105.
Dosdall, L., Lehmkuhl, D. M. 1979. Stoneflies (Plecoptera) of
Saskatchewan. Quaestiones Entomologicae 15:3 – 116.
Downing, J. A., Rigler, F. H. 1984. A manual on methods for the assessment of secondary productivity in fresh waters. 2nd ed. IBP
Handbook 17, Blackwell Scientific, Oxford, UK, xxiv 501 pp.
Drake, C. J., Harris, H. M. 1934. The Gerrinae of the Western
Hemisphere (Hemiptera). Annals of the Carnegie Museum
23:179 – 240.
Dunkle, S. W. 1977. Larvae of the genus Gomphaeschna (Odonata:
Aeshnidae). Florida Entomologist 60:223 – 225.
Dunn, C. E. 1979. A revision and phylogenetic study of the genus
Hesperocorixa Kirkaldy (Hemiptera: Corixidae). Proceedings of
the Academy of Natural Sciences of Philadelphia 131:158 – 190.
Edmunds, G. F., Jr. 1972. Biogeography and evolution of
Ephemeroptera. Annual Review of Entomology 17:21 – 42.
Edmunds, G. F., Jr., Jensen, S. L., Berner, L. 1976. The mayflies of
North and Central America. Uni. of Minnesota Press, Minneapolis, x 330 pp.
Edwards, S. W. 1961. The immature stages of Xiphocentron mexico
(Trichoptera). Texas Journal of Science 13:51 – 56.
Edwards, S. W., Arnold, C. R. 1961. The caddisflies of the San Marcos River. Texas Journal of Science 13:398 – 415.
Epler, J. H. 1987. Revision of the Nearctic Dicrotendipes Kieffer,
1913 (Diptera: Chironomidae). Evolutionary Monographs 9,
102 pp. 37 pl.
Epler, J. H. 1995. Identification manual for the larval Chironomidae
(Diptera) of Florida. Florida Department of Environmental Protection, Tallahassee. 317 pp.
Epler, J. H. 1996. Identification manual for the water beetles of
Florida. Florida Department of Environmental Protection, Tallahassee, iv 253 pp.
Exner, K., Craig, D. A. 1976. Larvae of Alberta Tanyderidae (Diptera:
Nematocera). Quaestiones Entomologicae 12:219 – 237.
Fall, H. C. 1922. A review of the North American species of Agabus
together with a description of a new genus and species of the
tribe Agabini. John D. Sherman, Jr., Mt. Vernon, NY, 36 pp.
Fall, H. C. 1923. A revision of the North American species of
Hydroporus and Agaporus. John D. Sherman, Jr., Mt. Vernon,
NY, 238 pp.
Ferkinhoff, W. D., Gundersen, R. W. 1983. A key to the whirligig
beetles of Minnesota and adjacent states and Canadian
provinces (Coleoptera: Gyrinidae). Scientific Publications of the
Science Museum of Minnesota, New Series 5(3):1 – 55.
Flint, O. S., Jr. 1960. Taxonomy and biology of Nearctic limnephelid
larvae (Trichoptera), with special reference to species in eastern
United States. Entomologica Americana 40:1 – 120.
724
W. L. Hilsenhoff
Flint, O. S., Jr. 1961. The immature stages of the Arctopsychinae occurring in eastern North America (Trichoptera: Hydropsychidae).
Annals of the Entomological Society of America 54:5 – 11.
Flint, O. S., Jr. 1962. Larvae of the caddisfly genus Rhyacophila in
eastern North America (Trichoptera: Rhyacophilidae). Proceedings of the United States National Museum 113:465 – 493
Flint, O. S., Jr. 1964. Note on some Nearctic Psychomyiidae with
special reference to their larvae (Trichoptera). Proceedings of
the United States National Museum 115:467 – 481.
Flint, O. S., Jr. 1984. The genus Brachycentrus in North America, with
a proposed phylogeny of the genera of Brachycentridae (Trichoptera). Smithsonian Contributions to Zoology 398, 58 pp.
Flowers, R. W. 1982. Review of the genus Macdunnoa
(Ephemeroptera: Heptageniidae) with description of a new
species from Florida. Great Lakes Entomologist 15:25 – 30.
Flowers, R. W., Hilsenhoff, W. L. 1975. Heptageniidae
(Ephemeroptera) of Wisconsin. Great Lakes Entomologist
8:201 – 218.
Floyd, M. A. 1995. Larvae of the caddisfly genus Oecetis (Trichoptera: Leptoceridae) in North America. Bulletin of the Ohio
Biological Survey New Series 10(3), viii 85 pp.
Foote, B. A. 1959. Biology and life history of the snail-killing flies belonging to the genus Sciomyza Fallen (Diptera: Sciomyzidae).
Annals of the Entomological Society of America 52:31 – 43.
Foote, B. A. 1961. Biology and immature stages of the snail-killing
flies belonging to the genus Tetanocera (Diptera: Sciomyzidae).
Ph.D. Dissertation, Cornell University, Ithaca, NY, 190 pp.
Foote, B. A. 1995. Biology of shore flies. Annual Review of Entomology 40:417 – 442.
Frania, H. E., Wiggins, G. B. 1995. Analysis of morphological and
behavioral evidence for the phylogeny and higher classification
of Trichoptera. Royal Ontario Museum Life Science Contribution 160.
Froeschner, R. C. 1949. Contribution to a synopsis of the Hemiptera
of Missouri. Part IV. Hebridae, Mesoveliidae, Cimicidae, Anthocoridae, Cryptostemmatidae, Isometopidae, Miridae. American Midland Naturalist 42:123 – 188.
Froeschner, R. C. 1962. Contribution to a synopsis of the Hemiptera
of Missouri. Part V. Hydrometridae, Gerridae, Veliidae, Saldidae, Ochteridae, Gelastocoridae, Naucoridae, Belostomatidae,
Nepidae, Notonectidae, Pleidae, Corixidae. American Midland
Naturalist 67:208 – 240.
Fullington, K. E., Stewart, K. W. 1980. Nymphs of the stonefly genus
Taeniopteryx (Plecoptera: Taeniopterygidae) of North America.
Journal of the Kansas Entomological Society 53:237 – 259.
Funk, D. H., Sweeney, B. W. 1994. The larvae of eastern North
American Eurylophella Tiensuu (Ephemeroptera: Ephemerellidae). Transactions of the American Entomological Society
120:209 – 286.
Garrison, R. W. 1981. Description of the larva of Ischnura gemina
with a key and new characters for the separation of sympatric
Ischnura larvae. Annals of the Entomological Society of America 56:356 – 364.
Garrison, R. W. 1984. Revision of the genus Enallagma of the United
States west of the Rocky Mountains and identification of certain larvae by discriminant analysis (Odonata: Coenagrionidae).
Uni. of California Publications in Entomology Vol. 105, ix
129 pp.
Gentili, E. 1985. I Laccobius americani - I Laccobius del Canada.
Osservatorio di Fisica Terrestre e Museo Antonio Stoppani del
Seminario Arcivescovile di Milano 6 (1983):31 – 45.
Gentili, E. 1986. I Laccobius americani - II genere Laccobius a sud del
Canada. Osservatorio di Fisica Terrestre e Museo Antonio Stoppani del Seminario Arcivescovile di Milano 7 (1984):31 – 40.
Gillespie, J. M., Barr, W. F., Elliott, S. T. 1994. Taxonomy and biology of the immature stages of species of Thaumalea occurring
in Idaho and California (Diptera: Thaumaleidae). Myia
5:153 – 193.
Givens, D. R., Smith, S. D. 1980. A synopsis of the western Arctopsychinae (Trichoptera: Hydropsychidae). Melanderia 35:1 – 24.
Glover, J. B. 1996. Larvae of the caddisfly genera Triaenodes and
Ylodes (Trichoptera: Leptoceridae) in North America. Bulletin
of the Ohio Biological Survey New Series 11(2), vii 89 pp.
Glukhova, V. M. 1977. Ceratopogonidae. Pages 431 – 457, in The
identification of fresh water invertebrates of European USSR i.e.
USSR west of the Urals (plankton and benthos). Zoological Institute., USSR Academy of Science. (translated from the Russian
by D.S. Kettle).
Gonsoulin, G. J. 1973a. Seven families of aquatic and semiaquatic
Hemiptera in Louisiana. Part I. Family Hydrometridae Billberg,
1820. “marsh treaders”. Entomological News 84:9 – 16.
Gonsoulin, G. J. 1973b. Seven families of aquatic and semiaquatic
Hemiptera in Louisiana. Part II. Family Naucoridae Fallen,
1814. “creeping water bugs”. Entomological News 84:83 – 88.
Gonsoulin, G. J. 1973c. Seven families of aquatic and semiaquatic
Hemiptera in Louisiana. Part III. Family Belostomatidae Leach,
1815. Entomological News 84:173 – 189.
Gonsoulin, G. J. 1974. Seven families of aquatic and semiaquatic
Hemiptera in Louisiana. Part IV. Family Gerridae. Transactions
of the American Entomological Society 100:513 – 546.
Gonsoulin, G. J. 1975. Seven families of aquatic and semiaquatic
Hemiptera in Louisiana. Part V. Family Nepidae Latreille, 1802.
Entomological News 86:23 – 32.
Goodwin, J. T. 1987. Immature stages of some eastern Nearctic Tabanidae (Diptera). VIII. Additional species of Tabanus Linnaeus
and Chrysops Meigen. Florida Entomologist 70:268 – 277.
Gordon, R. D. 1969. A revision of the niger-tenebrosus group of
Hydroporus (Coleoptera: Dytiscidae) in North America. Ph.D.
Dissertation, North Dakota State University, Fargo., 311 pp.
17 pl.
Gordon, R. D. 1981. New species of North American Hydroporus,
niger-tenebrosus group (Coleoptera: Dytiscidae). Pan-Pacific Entomologist 57:105 – 123.
Gordon, R. D., Post, R. L. 1965. North Dakota water beetles. North
Dakota Insects Publication No. 5. Department of Entomology,
North Dakota State University, Fargo., 52 pp.
Grodhaus, G. 1987. Endochironomus Kieffer, Tribelos Townes,
Synendotendipes, n. gen., and Endotribelos, n. gen. (Diptera:
Chironomidae) of the Nearctic region. Journal of the Kansas
Entomological Society 60:167 – 247.
Gundersen, R. W. 1978. Nearctic Enochrus. Biology, keys, descriptions and distribution (Coleoptera: Hydrophilidae). Department
of Biological Science, St. Cloud State University, MN., 54 pp.
Gundersen, R. W., Otremba, C. 1988. Haliplidae of Minnesota. Scientific Publications of the Science Museum of Minnesota, New
Series 6(3):1 – 43.
Haddock, J. D. 1977. The biosystematics of the caddisfly genus Nectopsyche in North America with emphasis on the aquatic stages.
American Midland Naturalist 98:382 – 421.
Hall, F. 1974. A key to the Simulium larvae of southern California
(Diptera: Simuliidae). California Vector Views 21:65 – 71.
Harden, P, Mickel, C. 1952. The stoneflies of Minnesota (Plecoptera). Technical Bulletin of the University of Minnesota
Agricultural Experiment Station 210:1 – 84.
Harper, P. P., Hynes, H. B. N. 1971a. The Leuctridae of eastern
Canada (Insecta; Plecoptera). Canadian Journal of Zoology
49:915 – 920.
Harper, P. P., Hynes, H. B. N. 1971b. The Capniidae of eastern
17. Diversity and Classification of Insects and Collembola
Canada (Insecta; Plecoptera). Canadian Journal of Zoology
49:921 – 940.
Harper, P. P., Hynes, H. B. N. 1971c. The nymphs of the Taeniopterygidae of eastern Canada (Insecta; Plecoptera). Canadian
Journal of Zoology 49:941 – 947.
Harper, P. P., Hynes, H. B. N. 1971d. The nymphs of the Nemouridae of eastern Canada (Insecta: Plecoptera). Canadian Journal
of Zoology 49:1129 – 1142.
Hatch, M. H. 1930. Records and new species of Coleoptera from
Oklahoma and western Arkansas, with subsidiary studies. Publications of the University of Oklahoma Biological Survey
2:15 – 26.
Hatch, M. H. 1953. The beetles of the Pacific Northwest. Part I: Introduction and Adephaga. University of Washington Publications in Biology 16, 340 pp.
Hatch, M. H. 1965. The beetles of the Pacific Northwest. Part IV:
Macrodactyles, Palpicornes, and Heteromera. Uni. of Washington Publications in Biology 16, 268 pp.
Hellman, J. L. 1975. A taxonomic revision of the genus Hydrochus
of North America, Central America and West Indies. Ph.D. Dissertation, University of Maryland, College Park, 441 pp.
Henry, B. C. 1993. A revision of Neochoroterpes (Ephemeroptera:
Leptophlebiidae) new status. Transactions of the American Entomological Society 199:317 – 333.
Henry, T. J., Froeschner, R. C. Eds. 1988. Catalog of the Heteroptera, or true bugs, of Canada and the continental United
States. E.J. Brill, Publishers, Leiden, Netherlands, New York,
NY, xx 958 pp.
Herring, J. L. 1951. The aquatic and semiaquatic Hemiptera of
northern Florida. Part IV. Classification of habitats and keys to
the species. Florida Entomologist 34:146 – 161.
Hilsenhoff, W. L. 1973. Notes on Dubiraphia (Coleoptera: Elmidae)
with descriptions of five new species. Annals of the Entomological Society of America 66:55 – 61.
Hilsenhoff, W. L. 1975. Notes on Nearctic Acilius (Dytiscidae), with
the description of a new species. Annals of the Entomological
Society of America 68:271 – 274.
Hilsenhoff, W. L. 1980. Coptotomus (Coleoptera: Dytiscidae) in eastern North America with descriptions of two new species. Transactions of the American Entomological Society 105:461 – 471.
Hilsenhoff, W. L. 1984a. Identification and distribution of Baetisca
nymphs (Ephemeroptera: Baetiscidae) in Wisconsin. Great
Lakes Entomologist 17:51 – 52.
Hilsenhoff, W. L. 1984b. Aquatic Hemiptera of Wisconsin. Great
Lakes Entomologist 17:29 – 50.
Hilsenhoff, W. L. 1985. The Brachycentridae of Wisconsin (Trichoptera). Great Lakes Entomologist 18:149 – 154.
Hilsenhoff, W. L. 1986. Semiaquatic Hemiptera of Wisconsin. Great
Lakes Entomologist 19:7 – 19.
Hilsenhoff, W. L. 1987. Effectiveness of bottle traps for collecting
Dytiscidae (Coleoptera). Coleopterists Bulletin 41:377 – 380.
Hilsenhoff, W. L. 1990. Gyrinidae of Wisconsin, with a key to adults
of both sexes and notes on distribution and habitat. Great
Lakes Entomologist 23:77 – 91.
Hilsenhoff, W. L. 1991. Comparison of bottle traps with a D-frame
net for collecting adults and larvae of Dytiscidae and Hydrophilidae (Coleoptera). Coleopterists Bulletin 45:143 – 146.
Hilsenhoff, W. L. 1992. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). I. Introduction, key to genera of adults, and distribution, habitat, life cycle, and identification of species of Agabetinae, Laccophilinae and Noteridae. Great Lakes Entomologist 25:57 – 69.
Hilsenhoff, W. L. 1993a. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). II. Distribution, habitat, life cycle, and identifica-
725
tion of species of Dytiscinae. Great Lakes Entomologist
26:35 – 53.
Hilsenhoff, W. L. 1993b. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). III. Distribution, habitat, life cycle, and identification of species of Colymbetinae, except Agabini. Great Lakes
Entomologist 26:121 – 136.
Hilsenhoff, W. L. 1993c. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). IV. Distribution, habitat, life cycle, and identification of species of Agabini (Colymbetinae). Great Lakes Entomologist 26:173 – 197.
Hilsenhoff, W. L. 1994. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). V. Distribution, habitat, life cycle, and identification of species of Hydroporinae, except Hydroporus Clairville
sensu lato. Great Lakes Entomologist 26:275 – 295.
Hilsenhoff, W. L. 1995a. Dytiscidae and Noteridae of Wisconsin
(Coleoptera). VI. Distribution, habitat, life cycle, and identification of species of Hydroporus Clairville sensu lato (Hydroporinae). Great Lakes Entomologist 28:1 – 23.
Hilsenhoff, W. L. 1995b Aquatic Hydrophilidae and Hydraenidae of
Wisconsin (Coleoptera). I. Introduction, key to genera of adults,
and distribution, habitat, life cycle, and identification of species
of Helophorus Fabricius, Hydrochus Leach, and Berosus Leach
(Hydrophilidae), and Hydraenidae. Great Lakes Entomologist
28:25 – 53.
Hilsenhoff, W. L. 1995c. Aquatic insects of Wisconsin. Keys to Wisconsin genera and notes on biology, habitat, distribution and
species. Publication 3 of the Natural History Museums Council,
University of Wisconsin-Madison, 79 pp.
Hilsenhoff, W. L. 1995d. Aquatic Hydrophilidae and Hydraenidae of
Wisconsin (Coleoptera). II. Distribution, habitat, life cycle and
identification of species of Hydrobiini and Hydrophilini (Hydrophilidae: Hydrophilinae). Great Lakes Entomologist
28:97 – 126.
Hilsenhoff, W. L., Billmyer, S. J. 1973. Perlodidae (Plecoptera) of
Wisconsin. Great Lakes Entomologist 6:1 – 14.
Hilsenhoff, W. L., Brigham, W. U. 1978. Crawling water beetles of
Wisconsin (Coleoptera: Haliplidae). Great Lakes Entomologist
11:11 – 22.
Hilsenhoff, W. L., Schmude, K. L. 1992. Riffle beetles (Coleoptera:
Dryopidae, Elmidae, Lutrochidae, Psephenidae) of Wisconsin
with notes on distribution, habitat, and identification. Great
Lakes Entomologist 25:191 – 213.
Hitchcock, S. W. 1974. Guide to the Insects of Connecticut. Part VII.
The Plecoptera or stoneflies of Connecticut. State Geological
and Natural History Survey of Connecticut Bulletin 107, vi
262 pp.
Hoffman, C. E. 1939. Morphology of the immature stages of some
northern Michigan Donaciini (Chrysomelidae: Coleoptera). Papers of the Michigan Academy of Science, Arts and Letters
25:243 – 290 10 plates.
Hogue, C. L., Georgian, T. 1986. Recent discoveries in the Blepharicera
tenuipes group, including descriptions of two new species from
Apalachia (Diptera: Blephariceridae). Contributions to the Science
and Natural History Museum, Los Angeles County 377:1 – 20.
Hubbard, M. D. 1994. The mayfly family Behningiidae
(Ephemeroptera: Ephemeroidea): Keys to the recent species with
a catalog of the family. Great Lakes Entomologist 27:161 – 168.
Huggins, D. G., Brigham, W. U. 1982. Odonata. in: Brigham, A. R.,
Brigham, W. U., Gnilka, A. Eds. 1981. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 4.1 – 4.100.
Hungerford, H. B. 1933. The genus Notonecta of the world (Notonectidae-Hemiptera). University of Kansas Science Bulletin
21:5 – 193.
726
W. L. Hilsenhoff
Hungerford, H. B. 1948. The Corixidae of the Western Hemisphere.
University of Kansas Science Bulletin 32:1 – 288; 408 – 827.
Hungerford, H. B. 1954. The genus Rheumatobates Bergroth
(Hemiptera-Gerridae). University of Kansas Science Bulletin
36:529 – 588.
Hungerford, H. B., Evans, N. W. 1934. The Hydrometridae of the
Hungarian National Museum and other studies in the family. Annales historico-naturales Musei Nationalis Hungarici 28:31 – 112.
Hutchinson, G. E. 1945. On the species of Notonecta (HemipteraHeteroptera) inhabiting New England. Transactions of the Connecticut Academy of Arts and Sciences 36:599 – 605.
Hynes, H. B. N. 1976. The biology of Plecoptera. Annual Review of
Entomology 21: 135 – 153.
Jansson, A. 1972. Systematic notes and new synonymy in the genus
Cenocorixa (Hemiptera: Corixidae). Canadian Entomologist
104:449 – 459.
Kavanaugh, D. H. 1986. A systematic review of amphizoid beetles
(Amphizoidae: Coleoptera) and their phylogenetic relationships
to other Adephaga. Proceedings of the California Academy of
Sciences 44:67 – 109.
Kettle, D. S. 1977. Biology and bionomics of bloodsucking ceratopogonids. Annual Review of Entomology 22:33 – 51.
Kittle, P. D. 1977. A revision of the genus Trepobates Uhler
(Hemiptera: Gerridae). Ph.D. Dissertation, University of
Arkansas, Fayetteville. 255 pp.
Kittle, P. D. 1980. The water striders (Hemiptera: Gerridae) of
Arkansas. Arkansas Academy of Science Proceedings 34:68 – 71.
Knutson, L. V., Berg, C. O. 1964. Biology and immature stages of
snail-killing flies: the genus Elgiva (Diptera: Sciomyzidae). Annals of the Entomological Society of America 57:173 – 192.
Kondratieff, B. C., Kirchner, R. F., Stewart, K. W. 1988. A review of
Perlinella Banks (Plecoptera: Perlidae). Annals of the Entomological Society of America 81:19 – 27.
Kondratieff, B. C., Kirchner, R. F., Voshell, J. R., Jr. 1981. Nymphs
of Diploperla (Plecoptera: Perlidae). Annals of the Entomological Society of America 74:428 – 430.
Kondratieff, B. C., Voshell, J. R., Jr. 1984. The North and Central
American species of Isonychia (Ephemeroptera: Oligoneuriidae). Transactions of the American Entomological Society
110:129 – 244.
Kuitert, L. C. 1942. Gerrinae in the University of Kansas Collections.
University of Kansas Science Bulletin 28:113 – 143.
LaRivers, I. 1948. A new species of Pelocoris from Nevada, with notes
on the genus in the United States (Hemiptera: Naucoridae). Annals of the Entomological Society of America 41:371 – 376.
LaRivers, I. 1951. A revision of the genus Ambrysus in the United
States. University of California Publications in Entomology
8:277 – 338.
LaRivers, I. 1954. Nevada Hydrophilidae (Coleoptera). The American Midland Naturalist 52:164 – 174.
Larson, D. J. 1975. The predaceous water beetles (Coleoptera: Dytiscidae) of Alberta: systematics, natural history and distribution.
quaestiones Entomologicae 11:245 – 498.
Larson, D. J. 1987. Revision of North American species of IIybius
Erichson (Coleoptera: Dytiscidae), with systematic notes on
Palaearctic species. Journal of the New York Entomological Society 95:341 – 413.
Larson, D. J. 1989. Revision of North American Agabus Leach
(Coleoptera: Dytiscidae): introduction, key to species groups,
and classification of the ambiguus-, tristis-, and arcticus-groups.
Canadian Entomologist 121:861 – 919.
Larson, D. J. 1991. Revision of North American Agabus Leach
(Coleoptera: Dytiscidae): elongatus-, zetterstedti-, and confinisgroups. Canadian Entomologist 123:1239 – 1317.
Larson, D. J. 1994. Revision of North American Agabus Leach
(Coleoptera: Dytiscidae): lutosus-, obsoletus-, and fuscipennisgroups. Canadian Entomologist 126:135 – 181.
Larson, D. J. 1996. Revision of North American Agabus Leach
(Coleoptera: Dytiscidae): The opacus-group. Canadian Entomologist 128:613 – 665.
Larson, D. J. 1997. Revision of North American Agabus Leach
(Coleoptera: Dytiscidae): The seriatus-group. Canadian Entomologist 129:105 – 149.
Larson, D. J., Alarie, Y., Roughley, R. E. 2001. Dytiscidae of Canada
and Alaska. NRC Research Press, Ottawa, Canada, about
850 pp.
Larson, D. J., Roughley, R. E. 1990. A review of the species of Liodessus Guignot of North America north of Mexico with the
description of a new species (Coleoptera: Dytiscidae). Journal of
the New York Entomological Society 98:233 – 245.
Larson, D. J., Wolfe, R. W. 1998. Revision of North American
Agabus (Coleoptera: Dytiscidae): the semivittatus-group. Canadian Entomologist 130:27 – 54.
Lawrence, J. F., Britton, E. B. 1991. Colcoptera (beetles), in: CSIRO,
Aquatic insects of Australia, 2nd edn. Melbourne Uni. Press,
Australia, pp. 543 – 683.
Leech, H. B. 1938. A study of the Pacific Coast species of Agabus
Leach, with a key to the Nearctic species. M.S. Thesis, University of California, Berkeley.
Leech, H. B., Chandler, H. P. 1956. Aquatic Coleoptera. in: Usinger,
R. L., Edr. Aquatic insects of California with keys to North
American genera and California species. Uni. of California
Press, Berkeley, CA, pp. 293 – 371.
Lehmkuhl, D. M., Anderson, N. H. 1971. Contributions to the
biology and taxonomy of the Paraleptophlebia of Oregon
(Ephemeroptera: Leptophlebiidae). Pan-Pacific Entomologist
47:85 – 93.
Louton, J. A. 1982. Lotic dragonfly (Anisoptera: Odonata) nymphs
of the southeastern United States: identification, distribution
and historical biogeography. Ph.D. Dissertation, University of
Tennessee, Knoxville, 357 pp.
Lugo-Ortiz, C. R., McCafferty, W. P. 1995. Taxonomy of the
North and Central American species of Camelobaetidius
(Ephemeroptera: Baetidae). Entomological News 106:178 – 192.
Lugo-Ortiz, C. R., McCafferty, W. P. Waltz, R. D. 1994. Contribution to the taxonomy of the Panamerican genus Fallceon
(Ephemeroptera: Baetidae). Journal of the New York Entomological Society 102:460 – 475.
Lugo-Ortiz, C. R., McCafferty, W. P. 1998. A new North American
genus of Baetidae (Ephemeroptera) and key to Baetis complex
genera. Entomological News 109:345 – 353.
Lugo-Ortiz, C. R., McCafferty, W. P., Waltz, R. D. 1999. Definition
and reorganization of the genus Pseudocloeon (Ephemeroptera:
Baetidae) with new species descriptions and combinations.
Transactions of the American Entomological Society 125:1 – 37.
Malcolm, S. E. 1971. The water beetles of Maine: including the families Gyrinidae, Haliplidae, Dytiscidae, Noteridae, and Hydrophilidae. Technical Bulletin 48, Life Sciences and Agricultural Experiment Station, University of Maine, Orono., 49 pp.
Maschwitz, D. E., Cook, E. F. 1999. Revision of the Nearctic species
of Polypedilum (Polypedilum) and Polypedilum (Urespedilum)
(Diptera: Chironomidae). Ohio Biological Survey Bulletin, New
Series 12(3), vii 136 pp.
Matta, J. F. 1974. The insects of Virginia: No. 8. The aquatic
Hydrophilidae of Virginia (Coleoptera: Polyphaga). Research
Division Bulletin 94, Virginia Polytechnic Institute and State
University, Blacksburg., iv 44 pp.
Matta, J. F. 1976. The insects of Virginia: No. 10. The Haliplidae of
17. Diversity and Classification of Insects and Collembola
Virginia (Coleoptera: Adephaga). Research Division Bulletin
109, Virginia Polytechnic Institute and State University, Blacksburg, vi 26 pp.
Matta, J. F., Wolfe, G. W. 1981. A revision of the subgenus Heterosternuta Strand of Hydroporus Clairville (Coleoptera: Dytiscidae). Pan-Pacific Entomologist 57:176 – 219.
McCafferty, W. P. 1975. The burrowing mayflies (Ephemeroptera:
Ephemeroidea) of the United States. Transactions of the American Entomological Society 101:447 – 504.
McCafferty, W. P. 1977. Biosystematics of Dannella and related subgenera of Ephemerella (Ephemeroptera: Ephemerellidae). Annals of the Entomological Society of America 70:881 – 889.
McCafferty, W. P. 1981. Aquatic entomology. The fishermen’s and
ecologists’ illustrated guide to insects and their relatives. Science
Books International, Boston, MA, xv 448 pp.
McCafferty, W. P. 1991. Toward a phylogenetic classification of the
Ephemeroptera (Insecta): a commentary on systematics. Annals
of the Entomological Society of America 84:343 – 360.
McCafferty, W. P., Provonsha, A. V. 1993. New species, subspecies,
and stage descriptions of Texas Baetidae (Ephemeroptera). Proceedings of the Entomological Society of Washington 95:59 – 69.
McCafferty, W. P., Waltz, R. D. 1995. Labiobaetis (Ephemeroptera:
Baetidae): New status, new North American species, and related
new genus. Entomological News 27:209 – 216.
McCafferty, W. P., Wigle, M. J., Waltz, R. D. 1994. Contributions to
the taxonomy and biology of (Acentrella turbida (McDunnough) (Ephemeroptera: Baetidae). Pan-Pacific Entomology
70:301 – 308.
McCorkle, D. V. 1965. (Subfamily Elophorinae). in: Hatch, M. H.,
Edr. The beetles of the Pacific Northwest. Part IV. Macrodactyles, Palpicornes, and Heteromera. Uni. of Washington
Publications in Biology 16, pp. 23 – 38.
McWilliams, K. L. 1969. A taxonomic revision of the North American species of the genus Thermonectus Dejean (Coleoptera:
Dytiscidae). Ph.D. Dissertation, Indiana University, Bloomington., 220 pp.
Menke, A. S., Ed. 1979a. The semiaquatic and aquatic Hemiptera of
California (Heteroptera: Hemiptera). Bulletin of the California
Insect Survey 21, xi 166 pp.
Menke, A. S., 1979b. Family Belostomatidae/giant water bugs, electric light bugs, toe biters, in: Menke, A. S., Ed. The semiaquatic
and aquatic Hemiptera of California (Heteroptera: Hemiptera).
Bulletin of the California Insect Survey Vol. 21, pp. 76 – 86.
Merritt, R. W., Cummins, K. W. Eds. 1996. An introduction to the
aquatic insects of North America. 3 ed. Kendall/Hunt,
Dubuque, IA., xiii 862 p.
Merritt, R. W., Ross, D. H., Peterson, B. V. 1978. Larval ecology of
some lower Michigan black flies (Diptera: Simuliidae) with keys
to the immature stages. Great Lakes Entomologist 11;177 – 208.
Michael, A. G., Matta, J. F. 1977. The insects of Virginia: No. 12.
The Dytiscidae of Virginia (Coleoptera: Adephaga) (subfamilies:
Laccophilinae, Colymbetinae, Dytiscinae, Hydaticinae and Cybistrinae). Research Division Bulletin 124, Virginia Polytechnic
Institute and State University, Blacksburg, vi 53 pp.
Miller, D. C. 1965. Hydrophilidae, except Elophorinae and
Sphaeridiinae. in: Hatch, M. H., Ed. The beetles of the Pacific
Northwest. Part IV. Macrodactyles, Palpicornes, and Heteromera. Uni. of Washington Publications in Biology 16, pp.
21 – 23, 38 – 46.
Miller, K. B. 1998. Revision of the Nearctic Liodessus affinis (Say
1823) group (Coleoptera: Dytiscidae, Hydroporinae, Bidessini).
Entomologica Scandinavica 29:281 – 314.
Miller, P. L. 1987. Dragonflies. (Naturalists’ handbooks 7). Cambridge Uni. Press, Cambridge, UK. 84 pp.
727
Morihara, D. K., McCafferty, W. P. 1979. The Baetis larvae of North
America (Ephemeroptera: Baetidae). Transactions of the American Entomological Society 105:139 – 221.
Morrissette, R. 1979. Les Gyrinidae (Coleoptera) du Quebec. Fabreries 5:51 – 58.
Moulton, J. K., Adler, P. H. 1995. Revision of the Simulium jenningsi
species-group (Diptera: Simuliidae). Transactions of the American Entomological Society 121:1 – 57.
Moulton, J. K., Adler, P. H. 1997. The genus Ectemnia (Diptera:
Simuliidae): taxonomy, polytene chromosomes, new species,
and phylogeny. Canadian Journal of Zoology 75:1896 – 1915.
Moulton, S. R. II, Stewart, K. W. 1997. A new species of and first
record of the caddisfly genus Cnodocentron Schmid (Trichoptera: Xiphocentronidae) north of Mexico. in: Holzenthal,
R. W., Flint, O. S. Jr., Eds. Proceedings 8th International Symposium on Trichoptera, Ohio Biological Survey, Columbus, pp
343 – 347.
Musser, R. J. 1962. Dragonfly nymphs of Utah (Odonata:
Anisoptera). Uni. of Utah Biological Series 12(6), viii 66 pp.
Nations, V. L. 1994. A phylogenetic analysis of the North American
species of Setodes (Trichoptera: Leptoceridae) with descriptions
of the larvae and key to their identification. M.S. Thesis, University of Alabama, Tuscaloosa, 72 pp.
Needham, J. G., Traver, J. R., Hsu, Y-C. 1935. The biology of
mayflies. Comstock Pub Co. Inc., New York, NY, xiv 759 pp.
Needham, J. G., Westfall, M. J. Jr. 1955. A manual of the dragonflies
of North America (Anisoptera). Uni. of California Press, Berkeley., xii 615 pp.
Needham, J. G., Westfall, M. J., Jr., May, M. L. 2000. Dragonflies
of North America. Scientific Publishers, Gainesville, FL.
xv 940 pp.
Neff, S. E., Berg, C. O. 1962. Biology and immature stages of Hoplodictya spinicornis and H. setosa (Diptera: Sciomyzidae). Transactions of the American Entomological Society 88:77 – 93.
Neff, S. E., Berg, C. O. 1966. Biology and immature stages of malacophagous Diptera of the genus Sepedon (Sciomyzidae). Bulletin
of the Agricultural Experiment Station, Virginia Polytechnic Institute, Blacksburg 566:5 – 113.
Neunzig, H. H. 1966. Larvae of the genus Nigronia Banks (Neuroptera: Corydalidae). Proceedings of the Entomological Society
of Washington 68:11 – 16.
Nimmo, A. P. 1987. The adult Arctopsychidae and Hydropsychidae
(Trichoptera) of Canada and adjacent United States. Quaestiones Entomologicae 23:1 – 189.
Oliver, D. R., Roussel, M. E. 1982. The larvae of Pagastia Oliver
(Diptera: Chironomidae) with descriptions of three Nearctic
species. Canadian Entomologist 114:849 – 854.
Oliver, D. R., Roussel, M. E. 1983. Redescription of Brillia Kieffer
(Diptera: Chironomidae) with descriptions of Nearctic species.
Canadian Entomologist 115:257 – 279.
Oygur, S., Wolfe, G. W. 1991. Classification, distribution, and phylogeny of North American (north of Mexico) species of Gyrinus
Muller (Coleoptera: Gyrinidae). Bulletin of the American Museum of Natural History, New York 207, 97 pp.
Parfin, S. I., Gurney, A. B. 1956. The spongilla-flies, with special reference to those of the Western Hemisphere (Sisyridae: Neuroptera). Proceedings of the United States National Museum
105:421 – 529.
Parker, C. R. 1998. A review of Goerita (Trichoptera: Goeridae),
with description of a new species. Insecta Mundi 12:227 – 238.
Parker, C. R., Wiggins, G. B. 1985. The Nearctic caddisfly genus
Hesperophylax Banks (Trichoptera: Limnephilidae). Canadian
Journal of Zoology 63:2433 – 2472.
Parker, C. R., Wiggins, G. B. 1987. Revision of the caddisfly genus
728
W. L. Hilsenhoff
Psilotreta (Trichoptera: Odontoceridae). Life Science Contributions Royal Ontario Museum 144, 55 pp.
Pechuman, L. L., Webb, D. W. Teskey, H. J. 1983. The Diptera or
true flies of Illinois. I. Tabanidae. Bulletin of the Illinois Natural
History Survey 33:1 – 122.
Peckarsky, B. L., Fraissinet, P., Penton, M. A., Conklin, D. J., Jr.
1990. Freshwater macroinvertebrates of northeastern North
America. Cornell Uni. Press, Ithaca, NY, xi 422 pp.
Perkins, P. D. 1980. Aquatic beetles of the family Hydraenidae in the
Western Hemisphere: classification, biogeography and inferred
phylogeny (Insects: Coleoptera). Quaestiones Entomologicae
16:1 – 554.
Pescador, M. L., Berner, L. 1981. The mayfly family Baetiscidae
(Ephemeroptera). Part II Biosystematics of the genus Baetisca.
Transactions of the American Entomological Society 107:
163 – 228.
Pescador, M. L., Peters, W. L. 1980. A revision of the genus Homoeoneuria (Ephemeroptera: Oligoneuriidae). Transactions of
the American Entomological Society 106:357 – 393.
Pescador, M. L., Rassmussen, A. K., Harris, S. C. 1995. Identification
manual for the caddisfly (Trichoptera) larvae of Florida. Florida
Department of Environmental Protection, Tallahassee, iv
132 pp.
Peterson, B. V. 1970. The Prosimulium of Canada and Alaska
(Diptera: Simuliidae). Memoirs of the Entomological Society of
Canada 69, 216 pp.
Peterson, B. V., Kondratieff, B. C. 1995. The black flies (Diptera:
Simuliidae) of Colorado: An annotated list with keys, illustrations and descriptions of three new species. Memoirs of the
American Entomological Society 42:1 – 121.
Pinder, L. C. V. 1986. Biology of freshwater Chironomidae. Annual
Review of Entomology 31:1 – 23.
Poirrier, M. A., Arceneaux, Y. M. 1972. Studies on southern Sisyridae
(spongillaflies) with a key to the third-instar larvae and additional
sponge-host records. American Midland Naturalist 88:455 – 458.
Polhemus, D. A. 1997. Systematics of the genus Rhagovelia Mayr
(Heteroptera: Veliidae) in the Western Hemisphere (exclusive of
the angustipes complex). Monographs Thomas Say Publications
in Entomology, ii 386 pp.
Polhemus, J. T., Chapman, H. C. 1979. Family Mesoveliidae/water
treaders. Pages 39 – 42, in Menke, A. S., Ed. The semiaquatic
and aquatic Hemiptera of California (Heteroptera: Hemiptera).
Bulletin of the California Insect Survey 21.
Porter, T. W. 1950. Taxonomy of the American Hebridae and the
natural history of selected species. Ph.D. Dissertation, University of Kansas, Lawrence, 185 pp. 12 pl.
Poulton, B. C., Stewart, K. W. 1991. The stoneflies of the Ozark and
Quachita Mountains (Plecoptera). Memoirs of the American
Entomological Society 38, 116 pp.
Pritchard, G. 1983. Biology of Tipulidae. Annual Review of Entomology 28:1 – 22.
Provonsha, A. V. 1990. A revision of the genus Caenis in North
America (Ephemeroptera: Caenidae). Transactions of the American Entomological Society 116:801 – 884.
Ray, D. H., Stark, B. P. 1981. The Nearctic species of Hydroperla
(Plecoptera: Perlodidae). Florida Entomologist 64:385 – 395.
Resh, V. H. 1976. The biology and immature stages of the caddisfly
genus Ceraclea in eastern North America (Trichoptera: Leptoceridae). Annals of the Entomological Society of America
69:1039 – 1061.
Ricker, W. E. 1943. Stoneflies of southwestern British Columbia. Indiana University Publications, Science Series 12, 145 pp.
Ricker, W. E. 1952. Systematic studies in Plecoptera. Indiana University Publications, Science Series 18, 200 pp.
Roback, S. S. 1985. The immature chironomids of the eastern United
States. VI. Pentaneurini - genus Ablabesmyia. Proceedings of the
Academy of Natural Sciences of Philadelphia 137:73 – 128.
Roback, S. S. 1986a. The immature chironomids of the eastern
United States. VII. Pentaneurini — genus Monopelopia new
record with redescription of the male adults and description of
some Neotropical material. Proceedings of the Academy of Natural Sciences of Philadelphia 138:350 – 365.
Roback, S. S. 1986b. The immature chironomids of the eastern
United States. VIII. Pentaneurini — genus Nilotanypus with the
description of a new species from Kansas. Proceedings of the
Academy of Natural Sciences of Philadelphia 138:443 – 465.
Roback, S. S. 1987. The immature chironomids of the eastern United
States. IX. Pentaneurini — genus Labrundinia with the description of some Neotropical material. Proceedings of the Academy
of Natural Sciences of Philadelphia 139:159 – 209.
Roberts, C. H. 1913. Critical notes on the species of Haliplidae of
America north of Mexico with descriptions of new species.
Journal of the New York Entomological Society 21:91 – 123.
Roemhild, G. 1976. Aquatic Heteroptera (ture bugs) of Montana.
Montana Agricultural Experiment Station Research Report
102, 70 pp.
Ross, H. H. 1944. The caddisflies or Trichoptera of Illinois. Bulletin
of the Illinois Natural History Survey 23:1 – 326.
Roughley, R. E. 1990. A systematic revision of species of Dytiscus
(Coleoptera: Dytiscidae). Part I. Classification based on adult
stage. Quaestiones Entomologicae 26:383 – 557.
Roughley, R. E., Pengelly, D. H. 1981. Classification, phylogeny, and
zoogeography of Hydaticus Leach (Coleoptera: Dytiscidae) of
North America. Quaestiones Entomologicae 17:249 – 309.
Saether, O. A. 1970. Nearctic and Palaearctic Chaoborus (Diptera:
Chaoboridae). Bulletin 174, Fisheries Research Board of
Canada, vii 57 pp.
Saether, O. A., Wang, X. 1995. Revision of the genus Paraphaenocladius Thienemann, 1924 of the world (Diptera: Chironomidae:
Orthocladiinae). Entomologica Scandinavica Supplement
48:3 – 69.
Sailer, R. I. 1948. The genus Trichocorixa (Corixidae, Hemiptera).
University of Kansas Science Bulletin 32:289 – 407.
Sanderson, M. W. 1982. Aquatic and semiaquatic Heteroptera, in:
Brigham, A. R., Brigham, W. U., Gnilka, A. Eds. The aquatic insects and oligochaetes of North and South Carolina. Midwest
Aquatic Enterprises, Mahomet, IL, pp. 6.1 – 6.94.
Schaefer, K. F., Drew, W. A. 1968. The aquatic and semiaquatic
Hemiptera of Oklahoma. Proceedings of the Oklahoma Academy of Science 47:125 – 134.
Schefter, P. W., Wiggins, G. B. 1986. A systematic study of the Nearctic larvae of the Hydropsyche morosa group (Trichoptera: Hydropsychidae). Life Sciences Miscellaneous Publication Royal
Ontario Museum, 94 pp.
Schmid, F. 1968. La famille des Arctopsychides. Memoires de la Societe Entomologique du Quebec 1, 84 pp
Schmude, K. L. 1992. Revision of the riffle beetle genus Stenelmis
(Coleoptera: Elmidae) in North America, with notes on bionomics. Dissertation, Ph.D. University of Wisconsin – Madison,
vi 388 pp.
Schmude, K. L. 1999. Riffle beetles in the genus Stenelmis (Coleoptera:
Elmidae) from warm springs in southern Nevada: new species,
new status, and a key. Entomological News 110:1 – 12.
Schmude, K. L., Hilsenhoff, W. L. 1986. Biology, ecology, larval taxonomy, and distribution of Hydropsychidae (Trichoptera) in
Wisconsin. Great Lakes Entomologist 19:123 – 145.
Schuster, G. A., Etnier, D. A. 1978. A manual for the identification of
the larvae of the caddisfly genera Hydropsyche Pictet and
17. Diversity and Classification of Insects and Collembola
Symphitopsyche Ulmer in eastern and central North America
(Trichoptera: Hydropsychidae). United States Environmental
Protection Agency Environmental Monitoring and Support Laboratory 600/ 4-78-060, xii 129 pp.
Scudder, G. G. E. 1965. The Notonectidae (Hemiptera) of British Columbia. Proceedings of the Entomological Society of British Columbia 62:38 – 41.
Scudder, G. G. E. 1971. The Gerridae (Hemiptera) of British Columbia. Journal of the Entomological Society of British Columbia
68:3 – 10.
Sherberger, F. F., Wallace, J. B. 1971. Larvae of the southeastern
species of Molanna. Journal of the Kansas Entomological Society 44:217 – 224.
Shirt, D. B., Angus, R. B. 1992. A revision of the Nearctic water beetles related to Potamonectes depressus (Fabricius) (Coleoptera:
Dytiscidae). Coleopterists Bulletin 46:109 – 141.
Simpson, K. W. 1982. A guide to basic taxonomic literature for the
genera of North American Chironomidae (Diptera) — adults,
pupae, and larvae. Bulletin of the New York State Museum 447,
43 pp.
Simpson, K. W., Bode, R. W., Albu, P. 1983. Keys for the genus
Cricotopus adapted from “Revision der Gattung Cricotopus
van der Wulp und ihrer Verwandten (Diptera, Chironomidae)”
by Hirvenoja, M. Bulletin 450 New York State Museum, v
133 pp.
Sites, R. W., Polhemus, J. T. 1994. The Nepidae (Hemiptera) of the
United States and Canada. Annals of the Entomological Society
of America 87:27 – 42.
Sites, R. W., Polhemus, J. T. 1995. The Pelocoris fauna of Texas.
Southwestern Naturalist 40:249 – 254.
Smetana, A. 1974. Revision of the genus Cymbiodyta Bed.
(Coleoptera: Hydrophilidae). Memoirs of the Entomological Society of Canada 93, iv 113 pp.
Smetana, A. 1980. Revision of the genus Hydrochara Berth.
(Coleoptera: Hydrophilidae). Memoirs of the Entomological Society of Canada 111, iv 100 pp.
Smetana, A. 1985. Revision of the subfamily Helophorinae of the
Nearctic region (Coleoptera: Hydrophilidae). Memoirs of the
Entomological Society of Canada 131. 154 pp.
Smetana, A. 1988. Review of the family Hydrophilidae of Canada
and Alaska. Memoirs of the Entomological Society of Canada
142, 316 pp.
Smith, C. L. 1980. A taxonomic revision of the genus Microvelia
Westwood (Heteroptera: Veliidae) of North America including
Mexico. Dissertation, Ph. D. University of Georgia, Athens,
388 pp.
Smith, C. L., Polhemus, J. T. 1978. The Veliidae (Heteroptera) of
America north of Mexico — keys and check list. Proceedings of
the Entomological Society of Washington 80:56-68.
Smith, R. F., Pritchard, A. E. 1956. Odonata. in: Usinger, R. L., Ed.
Aquatic insects of California with keys to North American genera and California species. Uni. of California Press, Berkeley,
pp. 106 – 153.
Snoddy, E. L., Noblet, R. 1976. Identification of the immature black
flies (Diptera: Simuliidae) of the southeastern United States with
some aspects of the adult role in transmission of Leucocytozoon
smithi to turkeys. South Carolina Agricultural Experiment Station Technical Bulletin 1057, 58 pp.
Soldan, T. 1984. A revision of the Caenidae with ocellar tubercles in
the nymphal stage (Ephemeroptera). Acta Universitatis
Carolinae — Biologica 1982 – 1984:289 – 362.
Spangler, P. J. 1960. A revision of the genus Tropisternus
(Coleoptera: Hydrophilidae). Dissertation, Ph. D. University of
Missouri, Columbia, 364 pp.
729
Spangler, P. J. 1973. A description of the larva of Celina angustata
Aube (Coleoptera: Dytiscidae). Journal of the Washington
Academy of Science 63:165 – 168.
Stark, B. P. 1983. A review of the genus Soliperla (Plecoptera: Peltoperlidae). Great Basin Naturalist 43:30 – 44.
Stark, B. P. 1986. The Nearctic species of Agnetina (Plecoptera: Perlidae). Journal of the Kansas Entomological Society 59:437 – 445.
Stark, B. P., Lago, P. K. 1983. Studies of Mississippi fishflies (Megaloptera: Corydalidae: Chauliodinae). Journal of the Kansas Entomological Society 56:356 – 364.
Stark, B. P., Ray, D. H. 1983. A revision of the genus Helopicus (Plecoptera: Perlodidae). Freshwater Invertebrate Biology 2:16 – 27.
Stark, B. P., Szczytko, S. W. 1981. Contributions to the systematics of
Paragnetina (Plecoptera: Perlidae). Journal of the Kansas Entomological Society 54:625 – 648.
Stark, B. P., Szczytko, S. W. 1988. A new Malirekus species from
eastern North America (Plecoptera: Perlodidae). Journal of the
Kansas Entomological Society. 61:195 – 199.
Stewart, K. W., Stanger, J. A. 1985. The nymphs, and a new species,
of North American Setvena Illies (Plecoptera: Perlodidae). PanPacific Entomologist 61:237 – 244.
Stewart, K. W., Stark, B. P. 1988. Nymphs of North American stonefly genera (Plecoptera). Thomas Say Foundation Vol. 12, xiii
460 pp.
Stewart, K. W., Stark, B. P., Huggins, T. G. 1976. The stoneflies (Plecoptera) of Louisiana. Great Basin Naturalist 36:366 – 384.
Stone, A., Jamnback, H. A. 1955. The black flies of New York State
(Diptera: Simuliidae). Bulletin of the New York State Museum
379. 144 pp.
Stone, A., Snoddy, E. L. 1969. The blackflies of Alabama (Diptera:
Simuliidae). Bulletin of the Alabama Agricultural Experiment
Station, Auburn University 390, 93 pp.
Stonedahl, G. M., Lattin, J. D. 1982. The Gerridae or water striders
of Oregon and Washington (Hemiptera: Heteroptera). Oregon
State University Agricultural Experiment Station Technical Bulletin 144, 36 pp.
Stonedahl, G. M., Lattin, J. D. 1986. The Corixidae of Oregon and
Washington (Hemiptera: Heteroptera). Oregon State University
Agricultural Experiment Station Technical Bulletin 150, 84 pp.
Stribling, J. B. 1986. Revision of Anchytarsus (Coleoptera: Dryopoidea) and a key to the new world genera of Ptilodactylidae.
Annals of the Entomological Society of America 79:219 – 234.
Surdick, R. F., Kim, K. C. 1976. Stoneflies (Plecoptera) of Pennsylvania, a synopsis. Bulletin 808 The Pennsylvania State University
Agricultural Experiment Station, University Park, 73 pp.
Szczytko, S. W., Stewart, K. W. 1977. The stoneflies (Plecoptera) of
Texas. Transactions of the American Entomological Society
103:327 – 378.
Szczytko, S. W., Stewart, K. W. 1979. The genus Isoperla (Plecoptera) of western North America; holomorphology and systematics, and a new stonefly genus Cascadoperla. Memoirs of
the American Entomological Society 32, i 120 pp.
Tai, L. D. D. 1967. Biosystematic study of Sympetrum (Odonata: Libellulidae). Dissertation, Ph. D. Purdue University, West
Lafayette, IN, xviii 234 pp.
Tarter, D. C., Little, M. L., Kirchner, R. F., Watkins, W. D., Farmer,
R. G., Steele, D. 1975. Distribution of pteronarcid stoneflies in
West Virginia (Insecta: Plecoptera). Proceedings of the West Virginia Academy of Science 47:79 – 85.
Tennessen, K. J. 1993. The larva of Progomphus bellei Knopf and Tennessen (Anisoptera: Gomphidae). Odonatologica 22:373 – 378.
Teskey, H. J. 1969. Larvae and pupae of some eastern North American Tabanidae (Diptera). Memoirs of the Entomological Society
of Canada 63, 147 pp.
730
W. L. Hilsenhoff
Teskey, H. J. 1983. A revision of the North American species of Atylotus (Diptera: Tabanidae) with keys to adult and immature
stages. Proceedings of the Entomological Society of Ontario
114:21 – 43.
Teskey, H. J., Burger, J. F. 1976. Further larvae and pupae of eastern
North American Tabanidae (Diptera). Canadian Entomologist
108:1085 – 1096.
Testa, S., III, Lago, P. K. 1994. The aquatic Hydrophilidae (Colcoptera) of Mississippi. Mississippi Agricultural and Forestry
Experiment Station, University of Mississippi Technical Bulletin
193, 71 pp.
Thorp, J. H., Covich, A. P., Eds. 1991. Ecology and classification of
North American freshwater invertebrates. Academic Press, San
Diego, CA, x 911 pp.
Tidwell, M. A. 1973. The Tabanidae (Diptera) of Louisiana. Tulane
Studies in Zoology and Botany 18:1 – 95.
Truxal, F. S. 1953. A revision of the genus Buenoa (Hemiptera: Notonectidae). University of Kansas Science Bulletin 35:1351 – 1517.
Unzicker, J. D., Carlson, P. H. 1982. Epemeroptera. in: Brigham, A.
R., Brigham, W. U. Gnilka, A. Eds. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 3.1 – 3.97.
Unzicker, J. D., McCaskill, V. H. 1982. Plecoptera. in: Brigham, A.
R., Brigham, W. U. Gnilka, A. Eds. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 5.1 – 5.50.
Unzicker, J. D., Resh, V. H. Morse, J. C. 1982. Trichoptera. in:
Brigham, A. R., Brigham, W. U. Gnilka, A. Eds. The aquatic insects and oligochaetes of North and South Carolina. Midwest
Aquatic Enterprises, Mahomet, IL, pp. 9.1 – 9.138.
Usinger, R. L., Ed. 1956. Aquatic insects of California with keys to
North American genera and California species. Uni. of California Press, Berkeley, ix 508 pp.
Valley, K. R., Berg, C. O. 1977. Biology, immature stages, and new
species of snail-killing Diptera of the genus Dictya (Sciomyzidae). Search 7(2), 44 pp.
Van Tassell, E. 1966. Taxonomy and biology of the subfamily Berosinae of North and Central America and the West Indies
(Coleoptera: Hydrophilidae). Dissertation, Ph. D. Catholic University, Washington, DC, v 329 pp.
Vineyard, R. N., Wiggins, G. B. 1988. Further revision of the caddisfly
family Uenoidae (Trichoptera): evidence for inclusion of Neophylacinae and Thremmatidae. Systematic Entomology 13:361 – 372.
Voigt, W. G., Garcia, R. 1976. Key to the Notonecta nymphs of the
West Coast United States (Hemiptera: Notonectidae). Pan-Pacific Entomologist 52:172 – 176.
Walker, E. M. 1953. The Odonata of Canada and Alaska. Vol. 1.
Part I: General. Part II: The Zygoptera — Damselflies. Uni. of
Toronto Press, Toronto, Ont, xi 292 pp.
Walker, E. M. 1958. The Odonata of Canada and Alaska. Vol. 2.
Part III: The Anisoptera — four families. Uni. of Toronto Press,
Toronto, Ont, xi 318 pp.
Walker, E. M., Corbet, P. S. 1975. The Odonata of Canada and
Alaska. Volume three. Part III: The Anisoptera — three families.
Uni. of Toronto Press, Toronto, Ont. xvi 308 pp.
Wallace, J. B., Merritt, R. W. 1980. Filter-feeding ecology of aquatic
insects. Annual Review of Entomology 25:103 – 132.
Wallace, J. B., Woodall, W. R. Staats, A. A. 1976. The larval
dwelling-tube, capture net and food of Phylocentropus placidus
(Trichoptera: Polycentropdidae). Annals of the Entomological
Society of America 69:149 – 154.
Wallis, J. B. 1933. Revision of the North American species, (north of
Mexico), of the genus Haliplus, Latreille. Transactions of the
Royal Canadian Institute 19:1 – 76.
Wallis, J. B. 1939. The genus Graphoderus Aube in North America
(north of Mexico) (Coleoptera). Canadian Entomologist
71:128 – 130.
Waltz, R. D., McCafferty, W. P. 1979. Freshwater springtails (Hexopoda: Collembola) of North America. Purdue University Agricultural Experiment Station, West Lafayette, IN, 32 pp.
Waltz, R. D., McCafferty, W. P. 1983. Austrotinodes Schmid (Trichoptera: Psychomyiidae), a first U.S. record from Texas. Proceedings of the Entomological Society of Washington 85:
181 – 182.
Waltz, R. D., McCafferty, W. P. 1987a. Revision of the genus
Cloeodes Traver (Ephemeroptera: Baetidae). Annals of the Entomological Society of America 80:191 – 207.
Waltz, R. D., McCafferty, W. P. 1987b. New genera of Baetidae for
some Nearctic species previously included in Baetis Leach
(Ephemeroptera). Annals of the Entomological Society of America 80:667 – 670.
Waltz, R. D., McCafferty, W. P. 1987c. Systematics of Pseudocloeon,
Acentrella,
Baetiella,
and
Liebebiella,
new
genus
(Ephemeroptera: Baetidae). Journal of the New York Entomological Society 95:553 – 568.
Waltz, R. D., McCafferty, W. P. 1989. New species, redescriptions, and
cladistics of the genus Pseudocentroptiloides (Ephemeroptera:
Baetidae). Journal of the New York Entomological Society
97:151 – 158.
Waltz, R. D., McCafferty, W. P. 1999. Additions to the taxonomy of
Americabaetis (Ephemeroptera: Baetidae): A. lugoi, n. sp., adult
of A. robacki, and key to larvae. Entomological News
110:39 – 44.
Waters, T. F. 1972. The drift of stream insects. Annual Review of Entomology 17:253 – 272.
Weaver, J. S. III. 1988. A synopsis of the North American Lepidostomatidae (Trichoptera). Contributions of the American Entomological Institute 24(2), iv 141 pp.
Weaver, J. S. III, Morse, J. C. 1986. Evolution of feeding and casemaking behavior in Trichoptera. Journal of the North American
Benthological Society 5:150 – 158.
Weaver, J. S., Sykora, J. L. 1979. The Rhyacophila of Pennsylvania,
with larval descriptions of R. banksi and R. carpenteri (Trichoptera: Rhyacophilidae). Annals of the Carnegie Museum
48:403 – 423.
Webb, D. W. 1994. The immature stages of Suragina concinna
(Williston) (Diptera: Athericidae). Journal of the Kansas Entomological Society 67:421 – 425.
Webb, D. W., Brigham, W. U. 1982. Aquatic Diptera. in: Brigham, A.
R., Brigham, W. U. Gnilka, A. Eds. The aquatic insects and
oligochaetes of North and South Carolina. Midwest Aquatic
Enterprises, Mahomet, IL, pp. 11.1 – 11.111.
Westfall, M. J., May, M. L. 1996. Damselflies of North America. Scientific Publications, Gainesville, FL, 650 pp.
Westfall, M. J., Jr., Tennessen, K. J. 1979. Taxonomic clarification
within the genus Dromogomphus Selys (Odonata: Gomphidae).
Florida Entomologist 62:266 – 273.
White, D. S. 1978. A revision of the Nearctic Optioservus
(Coleoptera: Elmidae), with descriptions of new species. Systematic Entomology 3:59 – 74.
Wiederholm, T., Ed. 1983. Chironomidae of the Holarctic region.
Keys and diagnosis. Part I. Larvae. Entomologica Scandinavica
Supplement 19, 457 pp.
Wiersema, N. A. 1998. Camelobaetidius variabilis (Ephemeroptera:
Baetidae), a new species from Texas, Oklahoma and Mexico.
Entomological News 109:21 – 26.
Wiersema, N. A., McCafferty, W. P. 1998. A new species of Pseudocentroptiloides (Ephemeroptera: Baetidae), with revisions to
other previously unnamed baetid species from Texas. Entomological News 109:110 – 116.
17. Diversity and Classification of Insects and Collembola
Wiggins, G. B. 1960. A preliminary systematic study of the North
American larvae of the caddisfly family Phryganeidae (Trichoptera). Canadian Journal of Zoology 38:1153 – 1170.
Wiggins, G. B. 1973. New systematic data for the North American
caddisfly genera Lepania, Goeracea and Goerita (Trichoptera:
Limnephilidae). Life Science Contributions Royal Ontario Museum 91, 33 pp.
Wiggins, G. B. 1977. Larvae of the North American caddisfly genera
(Trichoptera). Uni. of Toronto Press, Toronto, Ont, xi 401 pp.
Wiggins, G. B. 1981. Considerations on the relevance of immature
stages to the systematics of Trichoptera. in: Moretti, G. P., Ed.
Proceeding of the 3rd International Symposium on Trichoptera.
Series Entomology 20. Junk, Dr. W. The Hague, Netherlands,
pp. 395 – 407.
Wiggins, B. B. 1996. Larvae of the North American caddisfly genera
(Trichoptera). 2nd ed. Uni. of Toronto Press, Toronto, Ont,
Buffalo, NY, xiii 457 pp.
Wiggins, G. B., Richardson, J. S. 1982. Revision and synopsis of the
caddisfly genus Dicosmoecus (Trichoptera: Limnephilidae;
Dicosmoecinae). Aquatic Insects 4:181 – 217.
Wiggins, G. B., Wichard, W. 1989. Phylogeny of pupation in Trichoptera, with proposals on the origin and higher classification
of the order. Journal of the North American Benthological Society 8:260 – 276.
Wiggins, G. B., Wisseman, R. W. 1990. Revision of the North American caddisfly genus Desmona Trichoptera: Limnephilidae). Annals of the Entomological Society of America 82:155 – 161.
Willson, R. B. 1967. The Hydrophilidae of Michigan with keys to
species of the Great Lakes Region. Thesis, M. S. Michigan State
University, East Lansing, 100 pp.
Wilson, C. A. 1958. Aquatic and semiaquatic Hemiptera of Mississippi. Tulane Studies in Zoology 6:116 – 170.
Winters, F. C. 1926. Notes on the Hydrobiini (ColeopteraHydrophilidae) of boreal America. Pan-Pacific Entomologist
3:49 – 58.
Wojtowicz, J. A. 1982. A review of the adults and larvae of the genus
Pycnopsyche (Trichoptera: Limnephilidae) with revision of the
Pycnopsyche scabripennis (Rambur) and Pycnopsyche lepida
(Hagen) complexes. Dissertation, Ph. D. University of Tennessee, 292 pp.
Wolfe, G. W. 1984. A revision of the Hydroporus vittatipennis
species group of the subgenus Neoporus (Coleoptera: Dytiscidae). Transactions of the American Entomological Society
110:389 – 434.
Wolfe, G. W., Matta, J. F. 1981. Notes on nomenclature and classification of Hydroporus subgenera with the description of a new
genus of Hydroporini (Coleoptera: Dytiscidae). Pan-Pacific Entomologist 57:149 – 175.
Wolfe, G. W., Roughley, R. E. 1990. A taxonomic, phylogenetic, and
zoogeographical analysis of Laccornis Gozis (Coleoptera: Dytiscidae) with the description of Laccornini, a new tribe of Hydroporinae. Quaestiones Entomologicae 26:273 – 354.
Wood, D. M. 1978. Taxonomy of the Nearctic species of Twinnia and
Gymnopais (Diptera: Simuliidae) and a discussion of the ancestry of the Simuliidae. Canadian Entomologist 110:1297 – 1337.
Wood, D. M., Dang, P. T., Ellis, R. A. 1979. The insects and arachnids of Canada. Part 6. The mosquitoes of Canada (Diptera:
Culicidae). Agriculture Canada Publication 1686. 390 pp.
Wood, D. M., Peterson, B. V., Davies, D. M., Gyorkos, H. 1963. The
black flies (Diptera: Simuliidae) of Ontario. Part II. Larval
identification, with descriptions and illustrations. Proceedings of
the Entomological Society of Ontario 93:99 – 129.
Wood, F. E. 1962. A synopsis of the genus Dineutus (Coleoptera:
Gyrinidae) in the Western Hemisphere. M. S. Thesis, University
of Missouri, Columbia, 99 pp.
731
Wooldridge, D. P. 1966. Notes on Nearctic Paracymus with descriptions of new species (Coleoptera: Hydrophilidae). Journal of the
Kansas Entomological Society 39:712 – 725.
Wooldridge, D. P. 1967. The aquatic Hydrophilidae of Illinois. Illinois State Academy of Science 60:422 – 431.
Yamamoto, T. Wiggins, G. B. 1964. A comparative study of the North
American species in the caddisfly genus Mystacides (Trichoptera:
Leptoceridae). Canadian Journal of Zoology 42:1105 – 1126.
Young, F. N. 1953. Two new species of Matus, with a key to the
known species and subspecies of the genus (Coleoptera: Dytiscidae). Annals of the Entomological Society of America 46:49 – 55.
Young, F. N. 1954. The water beetles of Florida. Uni. of Florida
Press, Gainesville, FL, ix 238 pp.
Young, F. N. 1956. A preliminary key to the species of Hydrovatus
of the eastern United States (Coleoptera: Dytiscidae). Coleopterists Bulletin 10:53 – 54.
Young, F. N. 1963a. The Nearctic species of Copelatus Erichson
(Coleoptera: Dytiscidae). Quarterly Journal of the Florida
Academy of Science 26:56 – 77.
Young, F. N. 1963b. Two new North American species of Hydrovatus, with notes on other species (Coleoptera: Dytiscidae). Psyche
70:184 – 192.
Young, F. N. 1979a. A key to Nearctic species of Celina with descriptions of new species (Coleoptera: Dytiscidae). Journal of the
Kansas Entomological Society 52:820 – 830.
Young, F. N. 1979b. Water beetles of the genus Suphisellus Crotch in
the Americas north of Colombia (Coleoptera: Noteridae).
Southwestern Naturalist 24:409 – 429.
Young, F. N. 1981a. Predaceous water beetles of the genus
Desmopachria: the convexa-grana group (Coleoptera: Dytiscidae). Occasional Papers of the Florida Collection of Arthropods
2(III-IV):1 – 11.
Young, F. N. 1981b. Predaceous water beetles of the genus
Desmopachria Babington: the leechi-glabricula group
(Coleoptera: Dytiscidae). Pan-Pacific Entomologist 57:57 – 64.
Young, F. N. 1985. A key to the American species of Hydrocanthus
Say, with descriptions of new taxa (Coleoptera: Noteridae). Proceedings of the Academy of Natural Sciences, Philadelphia
137:90 – 98.
Young, W. C., Bayer, C. W. 1979. The dragonfly nymphs (Odonata:
Anisoptera) of the Guadelupe River Basin, Texas. Texas Journal
of Science 31:85 – 98.
Zalom, F. G. 1977. The Notonectidae (Hemiptera) of Arizona.
Southwestern Naturalist 22:327 – 336.
Zimmerman, J. R. 1970. A taxonomic revision of the aquatic beetle
genus Laccophilus (Dytiscidae) of North America. Memoirs of
the American Entomological Society 26., i 275 pp.
Zimmerman, J. R. 1981. A revision of the Colymbetes of North
America (Dytiscidae). Coleopterists Bulletin 35:1 – 52.
Zimmerman, J. R. 1985. A revision of the genus Oreodytes in North
America (Coleoptera: Dytiscidae). Proceedings of the Academy
of Natural Sciences, Philadelphia 137:99 – 127.
Zimmerman, J. R., Smith, A. H. 1975. A survey of the Deronectes
(Coleoptera: Dytiscidae) of Canada, the United States, and
northern Mexico. Transactions of the American Entomological
Society 101:651 – 722.
Zimmerman, J. R., Smith, R. L. 1975. The genus Rhantus
(Coleoptera: Dytiscidae) in North America. Part I. General
account of the species. Transactions of the American Entomological Society 101:33 – 123.
Zloty, J., Pritchard G. 1997. Larvae and adults of Ameletus mayflies
(Ephemeroptera: Ameletidae) from Alberta. Canadian Entomologist 129:251 – 289.
This Page Intentionally Left Blank
18
AQUATIC INSECT ECOLOGY
Anne E. Hershey
Gary A. Lamberti
Department of Biology
University of North Carolina – Greensboro
Greensboro, North Carolina 27402
Department of Biological Sciences
University of Notre Dame
Notre Dame, Indiana 46556
I. Introduction
II. Aquatic Insect Communities
A. Insect Taxonomic Diversity
B. Physical Constraints
on Aquatic Insects
C. Lentic Insect Communities
D. Lotic Insect Communities
E. Saline Habitats
F. Specialized Habitats
III. Aquatic Insect Life Histories
IV. Insect-Mediated Processes
A. Detritivory
I. INTRODUCTION
Aquatic insects are abundant in most freshwater
habitats and often exhibit high diversity. They play significant roles in freshwater ecosystems, including serving as food items for nearly the full range of vertebrate
and invertebrate predators found in aquatic systems.
The ecology of aquatic insects has been studied from
many perspectives, including species diversity, life-history constraints, community structure, predator – prey
interactions, detritivory, grazing, and implications for
nutrient dynamics. Considerable information is available on insect responses to a variety of environmental
conditions, including factors that operate on landscapelevel scales. Thus, they often are used as indicators of
water-quality conditions in both lentic and lotic systems. Longitudinal trends in insect functional feeding
groups are an important component of the River Continuum Concept (see Fig. 6 in Chapter 2), a major paradigm in the discipline of stream ecology. Although the
ecological literature on aquatic insects is extensive,
most of it has been produced within the last 20 years,
reflecting current interest in the topic. In spite of this
extensive body of work, the enormous diversity of
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
V.
VI.
VII.
VIII.
IX.
B. Grazing
C. Predator – Prey Interactions
D. Insects as Conduits for Energy Flow
in Aquatic Food Webs
Secondary Production
Effects of Land Use on Aquatic
Insect Communities
Role of Disturbance
Aquatic Insects in Biomonitoring Studies
Summary
Literature Cited
insects combined with their widespread distribution
dictates that much remains to be learned.
This chapter provides a brief overview of aquatic
insect communities in both lentic (standing water) and
lotic (flowing water) habitats and describes the role of
insects in ecological processes within these ecosystems.
The discussion is placed in the context of how physical
and life-history factors constrain the distribution and
abundance of aquatic insects, thereby altering communities and ecosystem function.
II. AQUATIC INSECT COMMUNITIES
A. Insect Taxonomic Diversity
Insects are the most species-rich and often the
most abundant group of substrate-dwelling macroinvertebrates, and have successfully invaded virtually all
aquatic habitats. Although there are many specialized
habitats, most aquatic insects are found either in lotic
or lentic habitats (Table I), and are constrained to one
or the other habitat by physiological adaptations
(see below). We will follow the classification used
733
734
A. E. Hershey and G. A. Lamberti
TABLE I List of aquatic and partially aquatic orders of insects, with estimated number of North American species, preferred
habitat, typical generation time, feeding mode, and examples of common genera.
Order
Collembola
Estimated no. of NA
species
Generation
time (yr)
Primary feeding
mode (FFG)
Examples of common NA
genera
Ephemeroptera
600
Water surface;
semiaquatic
Lotic (lentic)
Odonata
Anisoptera
300
Lentic lotic
1
Predators
Zygoptera
Orthoptera
120
40
Lentic lotic
Water margin;
semiaquatic
1
1
Predators
Shredders
Plecoptera
6 families
3 families
Heteroptera
330
250
410
Lotic (lentic)
Lotic (lentic)
Lentic lotic
1
1
1
Megaloptera
Neuroptera
Trichoptera
50
6
1350
Lotic (lentic)
Lotic lentic
Lotic (lentic)
Lepidoptera
Coleoptera
770
5000
Lentic (lotic)
Lentic lotic;
water margin
1
1
1 (some
or 1)
1
1 (some
1)
Shredders
Predators
Predators, collectorgatherers
Predators
Piercers
All major FFGs
Diptera
Tipulidae
Culicidae
Simuliidae
1500
175
150
Lotic
Lentic
Lotic
1
1
1
Chironomidae
2000
Lotic lentic
1
Shredders, predators
Collector filterers
Collector-filterers,
scrapers
All major FFGs
Others
3200
Lotic lentic
1
All major FFGs
Internal parasites
1
Parasites
Hymenoptera
40
Primary (and 2°)
habitat
60
1
Collector-gatherers
Podura, Sminthurides
1
Collector-gatherers,
collector-filterers,
scrapers
Baetis, Hexagenia
Aeshna, Gomphus,
Libellula
Argia, Lestes
Conocephalus
Shredders, scrapers
All major FFGs
Pteronarcys, Capnia
Acroneuria, Isoperla
Belostoma, Gerris,
Trichocorixa
Sialis, Corydalus
Sisyra
Rhyacophila, Hydropsyche,
Glossosoma, Neophylax
Petrophila
Hydroporus, Gyrinus,
Psephenus
Tipula, Limonia
Aedes, Anopheles, Culex
Prosimulium, Simulium
Chironomus, Polypedilum,
Tanytarsus
Chaoborus, Chrysops,
Ephydra
Apsilops, Trichogramma
(NA, North American; FFG, functional feeding group.)
by Hilsenhoff in this volume (see Chapter 17). The
orders Ephemeroptera (mayflies), Odonata (dragonflies and damselflies), Plecoptera (stoneflies), Trichoptera (caddisflies), and Megaloptera (fishflies and
alderflies) are considered to be strictly aquatic in their
immature stages. Other insect orders are partially
aquatic (aquatic group listed in parentheses), including
Collembola (semiaquatic springtails), Neuroptera
(spongillaflies), Heteroptera (aquatic and semiaquatic
bugs), Coleoptera (water beetles), Diptera (aquatic
flies and midges, especially suborder Nematocera but
some Brachycera), and Lepidoptera (aquatic caterpillars). All of these latter groups have important aquatic
representatives as immatures. The Coleoptera and
Heteroptera have groups with adult as well as larval
stages that are aquatic. Even several species of Orthoptera (grasshoppers) are found in association with
aquatic habitats (Cantrall and Brusven, 1996). These
aquatic insect orders are treated taxonomically in
Chapter 17.
Species diversity in many of the aquatic insect orders is very high. A pristine stream may contain well
over one hundred species of insects, including many
that are difficult or impossible to identify in the immature aquatic stages. Ponds, lakes, and wetlands can also
exhibit very high diversity, depending on water-quality
conditions, littoral development, and substrate characteristics. Due to taxonomic problems associated with
this high diversity, the role of aquatic insects in aquatic
ecosystems is often difficult to study. To minimize this
problem, aquatic insects have been categorized into a
smaller number of functional feeding groups (see Merritt and Cummins, 1996a, b), as discussed in more detail later in this chapter.
18. Aquatic Insect Ecology
735
B. Physical Constraints on Aquatic Insects
The physical environment of aquatic ecosystems exerts substantial control over the population abundances
and hence community composition of insects. Still, it is
remarkable that virtually every North American freshwater environment has been colonized by aquatic insects, ranging from alpine lakes to desert hot springs,
oligotrophic streams to sewage treatment lagoons, and
treeholes to the extreme depths of large lakes. Within
these habitats, physical factors of particular importance
to aquatic insects include: (1) dissolved oxygen concentration; (2) water temperature; (3) water chemistry;
(4) type of substrate; and (5) hydrodynamics.
1. Dissolved Oxygen
As aerobic organisms, all insects must obtain sufficient oxygen to drive their metabolic machinery (Eriksen et al., 1996). This presents a particular challenge
for aquatic insects because water, even when saturated,
contains much less oxygen than do terrestrial environments (a maximum of about 15 ppm oxygen in water
compared to over 200,000 ppm in the air). Dissolved
oxygen concentrations are highly variable over time
and space, and oxygen may be totally lacking from
some aquatic habitats (termed “anoxia”). Furthermore,
oxygen concentrations in lentic systems generally decrease with depth (Fig. 1A). Because the vast majority
of aquatic insects are substrate dwellers, this presents a
potential physiological constraint. Most cold-water
oligotrophic lakes remain oxygenated throughout the
open water season because solubility increases as water
temperature decreases. However, even in cold climates,
shallow ponds and wetlands can become anoxic due to
respiratory oxygen demands under ice where no opportunity exists for oxygen replenishment. Deeper oligotrophic lakes typically have sufficient oxygen stores to
meet under-ice metabolic demands. The likelihood of
under-ice anoxia increases with productivity due to
proportionally increasing respiratory requirements and
decreases with depth due to high oxygen storage in the
hypolimnion. Thus, seasonal and spatial variation in
oxygen concentration greatly restricts the types and diversity of insects found in aquatic environments.
No insect lacking access to an oxygen supply can
survive for long, but some can cope with short-term
anoxia. For example, the chironomid midge Chironomus and related Chironomini (Diptera: Chironomidae)
contain a hemoglobinlike pigment (hence the common
name “bloodworm”) that allows them to store oxygen
and thus survive temporary anoxia. The phantom
midge Chaoborus (Diptera: Chaoboridae) is also found
in anoxic sediments, but uses a different strategy than
do bloodworms. Chaoborus migrates vertically at night
FIGURE 1 Oxygen features in lakes and streams. (A) Oxygen and
temperature profiles in a temperate stratified lake. (B) Oxygen “sag”
in a river receiving a point discharge of municipal waste. BOD,
biological oxygen demand; COD, chemical oxygen demand; PPr, primary production.
toward the surface both to obtain oxygen and to hunt
for prey. During the day, it returns to the deep, dark
sediments to evade its own predators. Shallower environments (e.g., ponds and marshes) that experience
oxygen depletion contain a more diverse array of insects
because they can obtain oxygen in a myriad of ways.
Some insects, including many heteropterans, carry an
air store (“bubble”) on their body that they occasionally replenish by swimming to the water surface (Fig.
2A). Others maintain a semipermanent connection to
the atmosphere (“breathing tube”) as exemplified by
mosquito larvae (Diptera: Culicidae) and rat-tailed
maggots (Diptera: Syrphidae). Yet other insects, such as
some beetle larvae (Coleoptera) and a genus of mosquitoes (Fig. 2B), use specialized spiracles to pierce the vascular tissues of rooted aquatic plants and thus “steal”
their oxygen (White and Brigham, 1996).
736
A. E. Hershey and G. A. Lamberti
flowing water. For example, perlid stoneflies perform
“push-ups” when oxygen declines, presumably to move
more oxygenated water across their ventral thoracic
gills. Burrowing mayflies of the genus Hexagenia undulate their body to draw water, and thus food and oxygen, through their burrows.
2. Water Temperature
FIGURE 2 Respiratory structures of aquatic insects. (A) Ventral air
bubble of the backswimmer Neoplea (Heteroptera: Pleidae). (B)
Longitudinal section of postabdominal respiratory siphon of the
mosquito larva Taeniorhynchus (Diptera: Culicidae), used to pierce
plant air stores. (C) Abdominal filamentous tracheal gills of the
caddisfly Limnephilus (Trichoptera: Limnephilidae). [Drawings (A)
and (B) from Merritt and Cummins, 1996b; (C) from Wiggins, 1996.]
In most unimpacted streams, dissolved oxygen
concentrations generally are near saturation and thus
oxygen rarely limits insect diversity. We often see the
greatest diversity of aquatic insects in small streams,
due in part to the abundance of oxygen. However, insects requiring a connection to atmospheric air are typically in low numbers, because flow would quickly displace these taxa far downstream. Thus, most lotic
insects obtain their oxygen by diffusion through the
body wall, often aided by “tracheal gills,” which are
thinly sclerotized evaginations of the body wall designed to increase surface area (Fig. 2C). Most representatives of the insect orders that are wholly aquatic
(e.g., Ephemeroptera, Plecoptera, and Trichoptera) possess such tracheal gills in the larval, aquatic state and
achieve their highest diversity in flowing waters. Gills,
however, rely on abundant dissolved oxygen for diffusion into the body, and thus gilled larvae can experience stress at low oxygen concentrations. Such oxygen
“sags” can occur in larger rivers with low turbulence
(and therefore low reaeration), especially at night when
photosynthesis turns off or where municipal or industrial pollutants deplete oxygen due to high respiratory
demand downstream of their input (Fig. 1B). In these
cases, the aquatic insect fauna will more closely mimic
that found in oxygen-poor standing waters (many
dipterans, for example). Some behavioral adaptations
allow insects to “weather” periods of low oxygen in
Water temperature also imposes constraints on
aquatic insects. Temperature directly affects insects by
regulating their metabolic rates and thus development
from egg to adult, and indirectly by influencing such
things as fluid dynamics, gas saturation constants, and
primary production rates (Hauer and Hill, 1996). Fluctuation in water temperature is found both spatially
and temporally. For example, stratified lakes show
marked temperature gradients from top to bottom,
while rivers can show horizontal (bank to bank) and
longitudinal (source to estuary) gradients in temperature. Low- to midorder streams also may exhibit pronounced diel temperature fluctuations, especially on
sunny days. Shallow unshaded wetlands may exhibit
diel fluctuations of 20° (e.g., Hershey et al., 1995a).
Temporal fluctuations can range from subfreezing to
tepid in northern wetlands. Thus, temperature variation imposes fundamental constraints on the types of
insects that may be present in many aquatic habitats.
Some aquatic insects prefer a narrow range of temperature, usually cool water (cold “stenotherms”),
whereas others can tolerate a broader range of temperatures (“eurytherms”). Consequently, temperature fluctuation can limit the types of insects found within a
habitat. Any significant change in water temperature,
as may result from thermal pollution or climate
change, will likely alter the species composition of
aquatic insects. Considerable variation in temperature
tolerance occurs within the orders of aquatic insects,
and so temperature preferences should be examined on
a species-specific basis.
The upper lethal limit for all but the most specialized species is between 30 and 40°C (Pritchard, 1991;
Wallace and Anderson, 1996), although most temperate aquatic habitats do not approach such temperatures. However, thermally destabilized habitats, such as
cooling water reservoirs, have temperatures which may
approach this limit intermittently or seasonally, and
abundance and richness of most taxa are negatively affected (see Thorp and Diggins, 1982). Types of insects
that can tolerate, and even thrive, in hot water include
various beetles and true flies, although species diversity
declines with increasing temperature. For example, in a
survey of hot springs, Brock (1978) recorded 60 species
of Coleoptera at 30°C; at 45°C only two species were
found. At the other extreme, many species exhibit
18. Aquatic Insect Ecology
growth at temperatures only a few degrees above freezing in both lentic (e.g., Butler, 1982) and lotic systems
(e.g., Rosenberg et al., 1977).
Aquatic insects have several strategies for surviving
subfreezing conditions. The antifreeze compounds produced by some species prevent freezing of bodily fluids
to several degrees below 0°C (Danks, 1978; Duffy and
Liston, 1985). Other species actually tolerate freezing
of extracellular tissues, which provides some protection
against intracellular freezing (Danks, 1978; Zachariassen, 1979; Ward and Stanford, 1982). Cocoons and
diapausing eggs are important strategies to prevent ice
damage (see Wallace and Anderson, 1996). Because
water and snow cover provide some degree of thermal
buffering, even streams and ponds in the arctic do not
drop more than several degrees below freezing. In
north temperate regions, where terrestrial temperatures
may reach 35°C, stream temperatures typically do
not drop below 1°C in pools and 2°C in riffles
(Fig. 3; Hershey unpublished data). In arctic ponds that
freeze solid, some insect species overwinter several
times as larvae (Butler, 1982). Thus, aquatic insects
only need mechanisms that provide them with a few
degrees of thermal buffering. In some streams, avoidance of ice scour may be a more serious problem for
737
benthic insects. Prior to the formation of surface ice,
anchor ice attached to the stream bed may form at
night due to cooling associated with back radiation
(Barnes, 1928). As the stream warms during the day,
anchor ice lifts off the bottom, scouring the stream bed
and disturbing resident insects (Hynes, 1970).
Temperature, and its variation, can influence
aquatic insect diversity in any freshwater habitat. For
streams, the River Continuum Concept (Vannote et al.,
1980; see also below) predicts that biological diversity
should be highest in midorder (medium-sized) streams
because temperature variation is highest, thereby providing the most temperature “niches” for insects. The
literature on this topic, however, is mixed. Insect faunal
diversity can be high in cold streams, which are
thought to be the ancestral habitat for aquatic insects
(Hynes, 1970). For example, Anderson and Hanson
(1987) cataloged over 325 species of aquatic insects
from a cool Oregon stream. On the other hand, Morse
et al. (1983) found over 500 species of aquatic insects
in a warmer coastal South Carolina stream. From
wood snags in the warm Ogeechee River in Georgia,
Benke and Jacobi (1994) collected over 20 species of
mayflies alone, suggesting that aquatic insects can be
diverse at a variety of temperatures.
3. Water Chemistry
FIGURE 3
Pool and riffle temperature in the French River,
Minnesota, which is strongly influenced by ice. Although air
temperatures may drop as low as about 30°C, aquatic insects
experience temperatures that are only a few degrees below freezing.
Many aspects of water chemistry can restrict the
occurrence or abundance of aquatic insects, including
pH, salinity, and concentrations of specific ions or
elements. Generally, it is the extremes in any of these
parameters that result in change to aquatic insect communities, while levels around the “average” have less
direct impact. Low pH, as is found in acidified lakes
and streams of pH 5 (due to acid deposition, mine
drainage, organic acids, or poor buffering) can alter
community composition such that only acid-tolerant
taxa are found (Hynes, 1970). However, it is unclear
whether it is the low pH itself that adversely affects
insects or some related factor. For example, iron can
precipitate under acidic conditions with an associated
reduction in oxygen. Acidic waters can also leach metals from soils and rock, especially aluminum, that can
increase drift and possibly have toxic effects on aquatic
insects (Hall et al., 1987). Regardless, some aquatic
insects thrive in acid streams and lakes (examples cited
in Hynes, 1970). Salinity gradients that form in coastal
estuaries, along saline lakes, and even from runoff
after road salting, can affect insects, most of which
are salt-intolerant. However, some insects such as
brine flies (Diptera: Ephydridae) thrive in warm, saline
water, where they have few competitors. Many other
chemical features, such as calcium concentration and
total ionic strength, have potential importance to
738
A. E. Hershey and G. A. Lamberti
aquatic insects, but little is known about the requirements of specific taxa.
4. Substrate and Flow
At the scale of local habitat, substrate and hydrodynamics are probably the most important factors
determining the types and abundance of aquatic insects
present. Swift currents in streams, wave action along
shorelines, wind-generated turbulent mixing, and tidal
action all present significant hydrodynamic challenges
to insects. Most aquatic insects spend at least part, if
not most, of their life cycle associated with the substrate, and hydrodynamic forces interact strongly with
substrate type to produce the habitat experienced by the
insect fauna. Hydrodynamic and substrate influences on
aquatic insects are discussed in more detail in the sections below on lentic and lotic insect communities.
Cummins and Merritt (1996) recognized eight categories of aquatic insect habits, primarily based on the
substrate or habitat they occupy. The first four groups
remain associated with the substrate for most of their
lives. “Burrowers” and “sprawlers” inhabit fine sediments where they either tunnel into the sediments or remain on top of the sediments, respectively. “Climbers”
move along the stems of vascular macrophytes or pieces
of detritus. “Clingers” attach themselves to the surfaces
of substrates exposed to water movement, such as rocks
in streams or along wave-swept lake shores. The final
four groups spend part, or most, or their lives moving
through or on the water. “Swimmers” periodically propel themselves through the water to change location, although they mostly remain attached to a substrate.
TABLE II
“Divers” split their time among swimming to the surface, diving back through the water, and clinging to
submerged objects. “Skaters” are adapted for life on
top of the water surface, where they use hydrophobic
body parts to move over water. “Planktonic” forms
float or swim about in open water.
C. Lentic Insect Communities
Lentic environments, including lakes, ponds, and
various types of wetlands offer a broad array of
aquatic habitats that have been exploited by a wide variety of insects. As in any community, the insect species
that are present reflect the physical, chemical, and biotic characteristics of the ecosystem, each of which
places major physiological, morphological, and trophic
constraints on aquatic insects. Insects exhibit a variety
of adaptations to these habitat constraints, which combined have a large impact on the structure of lentic insect communities. In addition, numerous trophic and
interspecific factors interact with the physical and
chemical constraints to yield the dynamic and diverse
insect communities that are present in lentic systems.
1. Littoral Communities
Littoral areas of ponds and lakes are typically better oxygenated, structurally more complex, and afford
more abundant and diverse food resources than do
profundal sediments of lakes. All of these factors lead
to a high diversity of insects. Littoral communities generally have aquatic insect representatives from most of
the aquatic orders (Table II), including nearly all of the
Aquatic insect communities in lentic habitats
Habitat type
Habitat characteristics
Typical insect community
References
Littoral
High O2, macrophytes,
rock, sand, or silt
substrates
Merritt and Cummins,
1996a, b; Ward, 1992
Eutrophic
profundal
Low O2, soft mineral
and organic sediments
Ephemeroptera (esp. Siphlonuridae, Baetidae, Heptageniidae,
Caenidae, Leptophlebiidae), Odonata (representatives
from most families), Diptera (many families, but esp.
diverse Chironomidae), Trichoptera (esp. Phryganeidae,
Limnephilidae, Leptoceridae), Plecoptera (some Perlidae
and Perlodidae), Hemiptera (most aquatic families),
Megaloptera (few genera), Neuroptera (both aquatic
genera), Coleoptera (adults and larvae of many families),
Hymenoptera (parasitoids only)
Chironomidae (esp. Chironomus, Procladius)
Oligotrophic
profundal
Wetlands
High O2, soft mineral
sediments
Variable hydrology,
variable O2, organic
sediments, macrophytes
Chironomidae (esp. Tanytarsini and Orthocladiinae)
High diversity of most lentic orders. Diptera and Coleoptera
are especially well represented.
Hilsenhoff, 1966;
Jonasson, 1972;
Merritt and Cummins,
1996a, b
Saether,1979
Rosenberg and Danks,
1987; Batzer and
Wissinger, 1996;
Hershey et al., 1998
18. Aquatic Insect Ecology
739
morphological and trophic types as well as the full size
range of aquatic species. Large taxa such as dragonflies, damselflies, many of the lentic mayflies, aquatic
beetles and bugs, and caddisflies are important members of littoral zone communities, and typically are uncommon in profundal zones (Table II). Aquatic insects
of littoral zones also show considerable variation in
habitat (see Ward, 1992). Many of these species use the
macrophytes themselves as habitat, although others are
characteristic inhabitants of inorganic and organic sediments and rocks. Macrophyte habitats usually have
higher density and species diversity of insects than less
structurally complex habitats, including many of the
larger taxa mentioned above (Crowder and Cooper,
1982; Gilinsky, 1984; Hershey, 1985a). Many highly
mobile species, particularly members of the Coleoptera
and Heteroptera, swim in the water column (plankton),
or use the upper or lower surface of the air – water interface (epineuston and hyponeuston, respectively).
Gyrinid beetles, referred to as whirligig beetles because
they swim in whirling swarms on the water surface, are
especially well adapted for their habitat; they have dorsal and ventral compound eyes permitting them to see
above and below the water surface simultaneously. The
burrowing mayfly, Hexagenia, is often very abundant
in littoral and sublittoral sediments of lakes, but is particularly sensitive to anoxia (Beeton, 1961; Rasmussen,
1988). A few Plecoptera (stoneflies) are found in erosional lentic habitats (White and Brigham, 1996), reflecting the high oxygen requirements of members of
this order. Smaller species, especially representing the
dipteran family Chironomidae, are usually very diverse
and present in high densities (Thorp and Bergey, 1981;
Crowder and Cooper, 1982; Gilinski, 1984; Hershey,
1985a).
FIGURE 4 Feeding activity of Chironomus tentans returns P to the
water column. At high density this can be a significant source of P to
the water column. [From Gallepp, 1979.]
2. Profundal Communities
3. Wetland Communities
Profundal zones of lakes have quite different communities than do littoral zones (Table II). In both oligotrophic and eutrophic lakes, chironomids most often
dominate the insect fauna. However, in eutrophic lakes,
where the profundal zone is often anoxic during much
of the summer, Chironomus and Procladius may be the
only insect genera present (Hilsenhoff, 1966; Saether,
1979). Chironomus feeds on suspended particles by
pumping water into either U-shaped or J-shaped burrows, ingesting the particles that impinge on its silken
funnel at the opening of the burrow (Walshe, 1947).
Feces are deposited on the sediment surface forming
mounds. At high density, this feeding mechanism can
be important in returning phosphorus to the water column (Fig. 4; Gallepp, 1979). Procladius is a free-living
chironomid, most often characterized as a predator
(Kajak, 1980; Vodopich and Cowell, 1984), but it also
The various types of wetlands also afford a variety
of habitats for aquatic insects. Water level is highly
variable in wetlands, by definition, and many wetland
types are subjected to periods of desiccation. Desiccation provides a variety of challenges for aquatic insects,
and results in somewhat different communities than are
found in more permanent lentic ecosystems. Hydroperiod is likely, therefore, to be the most important factor
affecting wetland insect communities (Wiggins et al.,
1980; Batzer and Wissinger, 1996; Hershey et al.,
1998). Insect species in wetland habitats exhibit either
desiccation-resistance or drought-avoidance strategies
(Wiggins et al., 1980). These strategies do not fall
neatly along taxonomic lines. Desiccation resistant insects, especially many Diptera, and some odonates,
caddisflies, and beetles, deposit drought-resistant eggs
or lay eggs in plant stems, which then hatch to
ingests a variety of sediment dwelling diatoms and
desmids (Hershey, 1986). Although ubiquitous in lentic
habitats Procladius also is tolerant of low oxygen conditions. In oligotrophic lakes, the chironomid community in the profundal zone is more diverse, but still
depauperate compared to littoral zones. The presence
of Chironomus spp. versus Tanytarsus spp. has been
considered indicative of lake trophic condition (e.g.,
Brinkhurst, 1974). An elaborate key to lake typology
which recognizes several categories of trophic conditions has been developed based on the occurrence of
chironomid genera and species (Saether, 1979), and
lake classification has a long history based on chironomid assemblages (Brundin, 1949; Brinkhurst, 1974).
740
A. E. Hershey and G. A. Lamberti
repopulate the habitat following the return of hydric
conditions (Wiggins et al., 1980, reviewed by Batzer
and Wissinger, 1996). Insects that avoid desiccation
have several different strategies. Some migrate to more
permanent habitats to complete one or more generations. Wing polymorphism (i.e., winged and unwinged
individuals) associated with this strategy is not uncommon and is particularly well-studied in water striders
(Spence and Anderson, 1994) and some beetles (see
Wallace and Anderson, 1996). Other insects have long
flight periods as adults and thus will spend the drought
cycle in the terrestrial environment (Wiggins et al.,
1980). This strategy is also used by chironomids and
mosquitoes, as well as some dragonflies and limnephilid caddisflies (Wiggins et al., 1980; Wissinger,
1988). These general strategies interact with life history
(see below) such that wetland insect communities exhibit a succession following the drought cycle with
dominance shifting from mosquitoes and chironomids
to larger taxa such as beetles, odonates, heteropterans,
and caddisflies (Batzer and Wissinger, 1996). The extreme of this cycle is that prolonged drought results in
very low density of insects (Fig. 5; Hershey et al.,
1999). Under these conditions the wetland fauna is
characterized by high abundance of molluscs and annelids and low abundance and species richness of insects, but it becomes more and more dominated by a
rich insect fauna with prolonged inundation (Fig. 6;
Hershey et al., 1999).
D. Lotic Insect Communities
Lotic insect communities are functionally and
structurally quite different than lentic communities due
to the different physical and chemical challenges of the
lotic environment. At a local scale in stream ecosystems, substrate and current velocity are probably the
most important physical factors determining the community structure of aquatic insects. The stream substrate has obvious importance because the vast majority of stream insects spend most of their lives attached
to substrates. Lotic substrates can be broadly divided
into inorganic substrates (geologic material ranging
from silt to boulders) and organic substrates (fine organic particles up to logs). The particle size of inorganic matter has a large influence on insect community
FIGURE 5 Prolonged drought in Minnesota prairie wetlands in 1987 – 1990 resulted in communities with
significantly lower density of insects (measured in 1989 – 1990) than in wetter years of 1991 – 1993. Much of
this difference was due to Chironomidae.
18. Aquatic Insect Ecology
741
FIGURE 6 Patterns of insect and noninsect abundances in prairie wetlands over a hydrologic cycle.
Insects and cladocerans are abundant during the wet phase, but molluscs and copepods dominate
during drought. Because of their longer life cycles, the transition back to insects can be relatively long
following the return of hydric conditions. [From Hershey et al., 1999.]
structure. Coarser bed materials (e.g., gravel, cobble,
and boulders) generally provide more interstitial habitat for insects than do finer sediments (e.g., sand and
silt). For example, “water penny” beetles (family
Psephenidae), hellgrammite larvae (Megaloptera: Corydalidae), and perlid stoneflies (Plecoptera) frequently
are found in interstitial spaces on the undersides of
rocks. Sedentary filter-feeding insects require the space
between particles both to operate their filtration device
(e.g., nets, specialized body parts) and to allow water
flow to carry food items to them. Many case-building
caddisflies (Trichoptera) pupate in dense aggregations
on protected faces of cobbles and boulders where interstitial water flow carries the dissolved oxygen needed
for metamorphosis (Lamberti and Resh, 1979; Resh
et al., 1981), but they are more dispersed in the larval
stage. A lower diversity of insects is typically found in
fine sediments because the tight packing of particles restricts physical habitat and the trapping of detritus, and
can limit the availability of oxygen (Allan, 1995).
However, taxa adapted to such habitats (e.g., chironomid midges, oligochaete worms, certain mayflies) may
be abundant. For example, Soluk (1985) recorded
densities of chironomid midges exceeding 80,000 individuals / m2 in the sand substrate of an Alberta river.
Organic substrates such as large woody debris or leaf
packs often are “hot spots” for invertebrate activity because they provide both substrate and nutritional resources (Anderson and Sedell, 1979). In a survey of
Oregon streams, Dudley and Anderson (1982) found
56 invertebrate taxa that were closely associated with
wood, and another 129 taxa that were facultative associates of wood. The more specialized moss habitat typically supports a high density of chironomids and other
flies (e.g., moth flies, family Psychodidae) and a different assemblage of species than found on adjacent mineral substrates (Glime and Clemons, 1972; Suren and
Winterbourn, 1992).
Flow affects many aspects of insect biology, including body form, food acquisition, and movement. Taxa
found in the swift currents of streams usually are
streamlined (e.g., Baetis mayflies), dorsoventrally
compressed (e.g., heptageniid mayflies and psephenid
“water penny” beetles), possess suctorial disks (e.g.,
blepharicerid flies), use rock “ballast” in their cases to
resist the flow (e.g., Glossosoma caddisflies), or are
small enough to fit almost entirely within the sheltered
“boundary layer” (e.g., chironomid midges). Many insects take advantage of swift flow by allowing current to
transport food items to them. Examples include caddisflies of the family Hydropsychidae that construct silken
nets to capture suspended food particles, black flies
742
A. E. Hershey and G. A. Lamberti
FIGURE 8 Channel cross section of a stream during summer low
flow, showing position of hyporheic zone relative to surface water
and groundwater. [Modified slightly from Williams, 1993].
FIGURE 7 The larval blackfly is a common filter feeder in riffle
habitats of North American streams. The fan-shaped objects, or
cephalic fans, are a modification of the mouthparts which collect fine
particulate organic matter (FPOM) from the water column. The larva
orients itself in the current by forming a silk pad on the rock, then
attaches itself to the pad using tiny hooks on the posterior portion of
its abdomen. [Photograph courtesy of D. A. Craig.]
(Simuliidae) that filter food with specialized mouthparts
(Fig. 7), and Brachycentrus caddisflies that capture
particles on the setae (hairs) of their front legs (Wallace
and Merritt, 1980). Filter-feeding insects also can be
found along the wave-swept shores of lakes or on stable
substrates (e.g., wood debris) of large rivers, where similar hydrodynamic forces are experienced. Many insects
prefer low water movement, such as found in quiescent
stream pools and along stream margins. In these habitats, organic matter is deposited by gravity and thus can
be “gathered” and ingested. These taxa often are more
“wormlike” than swift-water forms.
Variation in flow (floods to desiccation) is the major cause of natural disturbance in streams (see section
below on disturbance) and is responsible for large, usually temporary, reductions in macroinvertebrate abundance and diversity (Lamberti et al., 1991). The hyporheic (i.e., interstitial) zone and the undersurfaces of
rocks provide refugia during flood events (Fig. 8). Dur-
ing drought, the deep hyporheic zone is especially important, but is usually accessible only to smaller invertebrates (typically about 1 – 5 mm in length). The spatial
extent of the hyporheic zone varies with local geomorphology. In bedrock-dominated reaches, it is virtually
nonexistent; whereas in the Flathead River (Montana),
the hyporheic zone may extend tens of meters vertically
and up to three kilometers laterally from the river, supporting relatively large stoneflies several millimeters in
length (Stanford and Ward, 1988). Finally, recolonization by egg-laying adults or by individuals drifting from
upstream (Table III) serves to reintroduce taxa that become locally extinct during drought, flood, or other disturbance events (Fisher et al., 1982; Smock, 1996).
1. Functional Feeding Groups
Rather that focusing on the species as the ecological
unit in streams, aquatic insect ecologists often consider
the insect community in terms of functional feeding
groups (Cummins, 1973, 1974; Cummins and Klug,
1979; Merritt and Cummins, 1996a). The functional
feeding group approach categorizes stream consumers
according to their mode of feeding, or functional role,
rather than taxonomic groups. From an ecosystem perspective, this approach reduces the number of groups to
be considered by 1 – 2 orders of magnitude compared to
the species approach. Thus, rather than studying dozens
or hundreds of specific consumers, a smaller number of
groups of organisms can be studied collectively from the
perspective of their function in the stream ecosystem.
This categorization also has utility in other aquatic
ecosystems, and has recently been applied to all North
American aquatic insects regardless of habitat (Merritt
and Cummins, 1996b).
18. Aquatic Insect Ecology
TABLE III
743
Mechanisms of insect recolonization of stream reaches
Mechanism
Time scales
Example
References
Downstream drift
Many insects
Waters, 1972
Many insects
Cairns, 1990
Upstream flight
Flight from other watersheds
Nocturnal, constant, episodic, dispersal
stages of some invertebrates
Nocturnal, constant, episodic, dispersal stages
of some invertebrates
Seasonal, depending on emergence period
Seasonal, depending on emergence period
Baetis mayflies
Many insects, Baetis mayflies
Upstream swimming
Seasonal
Leptophlebia cupida,
Paraleptophlebia mayflies
Upstream crawling
Daily
Dicosmoecus caddisflies,
Juga snails
Movement from hyporheic
refuge
Episodic, seasonal
Many early instar insects;
overwintering stages of many
insects in cold climates; many
residents of intermittent streams
Hershey et al., 1993
Wallace et al., 1986;
Schmidt et al., 1995
Hayden and Clifford,
1974; Bird and
Hynes, 1981
Hart and Resh, 1980;
L. M. Blair and G. A.
Lamberti,
unpublished data
Sedell et al., 1990;
Delucchi, 1989;
Hershey unpublished
data
Drift from tributaries
[From Hershey and Lamberti, 1998]
The functional feeding groups recognized by
Merritt and Cummins (1996a, b) are: (1) “scrapers”
( “grazers”), which remove and consume attached algae and associated periphytic material; (2) “shredders,”
which ingest coarse particulate organic matter (CPOM),
as decomposing leaf litter, living macrophyte tissue, or
dead wood; (3) “predators,” which eat living animals;
and (4) “collectors,” which consume decomposing fine
particulate organic matter (FPOM). The last group can
be further broken down into “collector-gatherers,”
which collect FPOM from the sediments, and “collector-filterers,” which collect FPOM from the water column (Fig. 9). Two other, less common functional feeding groups are the “macrophyte-piercers,” which pierce
the tissues of macroalgae and rooted hydrophytes, and
“parasites,” which develop on or in aquatic insects,
generally killing them. Because groups of insect species
can be studied collectively to unravel major avenues of
organic matter processing, the functional feeding group
approach greatly simplifies the study of insect function
in aquatic ecosystems. It also provides a strong basis for
comparative studies of the biomass and productivity of
consumer groups across ecosystems, whereas it is much
more difficult (and often less informative) to make such
comparisons on a species-by-species basis.
Although implementation of this approach has
been an important precedent for the development of
other major paradigms of aquatic ecology (e.g., river
continuum concept of Vannote et al., 1980), the functional feeding group concept is not without limitations.
First, assignment of individual organisms from aquatic
samples to a functional feeding group generally requires
identification of the organism to family (see Chapter 17
of this volume) or genus. This can be a very timeconsuming task, although a shortcut approach using an
illustrated key is effective for many taxa (see Merritt
and Cummins, 1996a). Second, feeding ecology can
vary within a genus, and even within a species depending on habitat or food availability. Thus, published
functional group designations for the generic level are
not always reliable (Mihuc, 1997). In fact, many
aquatic insects are known to change feeding habit with
growth and development (Chapman and Demory, 1963;
Rader and Ward, 1987). For example, the large limnephilid caddisfly Dicosmoecus gilvipes begins larval
life as a collector-gatherer, feeds primarily as a scraper
of periphyton during the middle instars, and finally becomes partly predaceous in the final instar (Gotceitas
and Clifford, 1983; Li and Gregory, 1989). Other insects are omnivorous throughout their lives. Filterfeeding caddisflies of the genus Hydropsyche will consume virtually anything that becomes entrapped in their
nets, including FPOM, living algae, and small animals
(Benke and Wallace, 1980), thus making them potential
members of several functional feeding groups.
Some orders of aquatic insects, or the aquatic
representatives within an order, perform mostly as a
single functional feeding group, whereas others show
great diversity in feeding (Table I). Generally, the more
primitive orders of aquatic insects show more limited
diversity in feeding mode. For example, aquatic collembolans are exclusively collector-gatherers, odonates
are strictly predaceous, and stoneflies (Plecoptera) are
either shredders or predators, generally depending on
family. Less speciose orders also tend to show less
feeding diversity, as one might expect. For example,
744
A. E. Hershey and G. A. Lamberti
FIGURE 9 Simplified food web for a hypothetical North American forested stream, showing
major energy inputs and use by different macroinvertebrate functional feeding groups [modified
from Allan, 1995, after Cummins, 1973]. Macroinvertebrate examples are: shredders
(Peltoperla stonefly, Tipula cranefly, Hydatophylax caddisfly); collector-filterers (Hydropsyche
caddisfly, Simulium blackfly); grazers (Glossosoma caddisfly, Juga snail); predators (Calineuria
stonefly, Orohermes hellgrammite). Broken lines indicate fate of fecal matter.
megalopterans are entirely predaceous, neuopterans
pierce either animals or plants, and aquatic hymenopterans are mostly parasites. In contrast, more derived orders, such as Trichoptera, Diptera, and
Coloptera, show considerable diversity in feeding
mode, in part due to their large number of aquatic
species. The Diptera and Trichoptera, in particular,
have been enormously successful in freshwater ecosystems. Chironomid midges alone may equal the species
diversity of all other insects combined in a typical
aquatic ecosystem. In any specific stream, it is usually
possible to find caddisflies represented in all functional
feeding groups, and often dominating the biomass of
those groups. Beetles possibly occupy the broadest
range of aquatic habitats, ranging from ice fields to
seashores, and can be very speciose.
2. The River Continuum Concept
The River Continuum Concept or “RCC” (Vannote
et al., 1980; see Fig. 6 in Chapter 2) is an important
paradigm in stream ecology and functional feeding
groups are a major component of that paradigm. The
RCC predicts that small, heavily shaded streams will
have high inputs of detritus (“allochthonous” CPOM)
from adjacent riparian zones. Due to this abundant
detritus, shredders will dominate the macroinvertebrate
community. Both collector-filterers and collectorgatherers also will be abundant because high-quality
FPOM will be produced as CPOM becomes fragmented. In medium-sized streams, light inputs increase
and thus benthic algal production will increase. The
shredders are replaced by scrapers, while collectors remain abundant. In large rivers, benthic algal production
and direct riparian inputs decline. Food resources for
macroinvertebrates are dominated by FPOM in suspension; collectors dominate the macroinvertebrate community. In all streams, predators comprise a small but
fairly stable proportion of the fauna. These predictions
were tested by Hawkins and Sedell (1981) for the
McKenzie River drainage in Oregon, which consisted of
streams ranging in size from the 1st- to 7th-order. They
found that functional feeding group distributions were
generally consistent with predictions of the RCC (Fig.
10). These longitudinal shifts were more pronounced
18. Aquatic Insect Ecology
745
in six small streams in Oregon, but that absence of
canopy increased the abundance of all functional feeding groups possibly because both autotrophic (on-site)
and heterotrophic (from upstream) inputs were received. Hence, insect consumers responded to the abundance and composition of the available food resources.
This notion of energy transfer via both autotrophic and
heterotrophic food webs is an underpinning of the lotic
ecosystem model of McIntire and Colby (1978), that
postulates a hierarchical organization to stream food
webs and a central role for aquatic insects.
3. Insect Drift
FIGURE 10
Longitudinal changes in relative abundance of
macroinvertebrate functional feeding groups at different seasons
within an Oregon, USA, river system. “Collectors” collectorgatherers; “Filterers” collector-filterers. Stream size increases along
the x-axis; DCC, Devilsclub Creek; MACK, Mack Creek; LOKC,
Lookout Creek; and MZER, McKenzie River. Note the general
conformance to predictions of the River Continuum Concept (see
Figure 2.6 in this volume). [Modified slightly from Hawkins and
Sedell, 1981.]
than were seasonal fluctuations in community structure
in the individual Oregon streams. Not all studies of
functional feeding groups have shown the same degree
of conformance to the RCC (e.g., Minshall et al., 1985;
Winterbourn et al., 1981). Some groups, such as shredders, predictably decline with increasing stream size
whereas other groups, such as scrapers, do not fit
tightly with RCC expectations. In evaluating conformance to the RCC, abundance and biomass have often
been considered as currency. However, some researchers
have argued that production rather than biomass is a
more appropriate currency as it is a better measure of
transfer of energy between trophic levels (Benke, 1993,
1998; Grubaugh et al., 1997). Large rivers, at least in
their natural state, likely have more complex energy inputs than implied by the RCC. Thorp and Delong
(1994) developed the riverine productivity model that
suggests that riparian inputs and benthic primary production are quite important to insect production in
large rivers, which is contrary to predictions of the
RCC. Their preliminary test of the model in the Ohio
River suggested that the shallow edges of large rivers
function similarly (and have similar insect communities)
as midorder streams, because they receive riparian litterfall and also have substantial periphyton production.
Hawkins et al. (1982) found that riparian canopy
highly influenced functional feeding group composition
“Drift” is the downstream transport of oranisms in
the current (usually expressed as number / per cubic meters), and it is a unique property of stream ecosystems.
Because it provides insects with a unidirectional transportation system, it has the potential to be important to
the distribution and ecology of stream insects. At any
given time, the proportion of fauna in the drift, compared to that on the substrate, is very small (0.01 – 0.5%;
Ulfstrand, 1968; Waters 1972). Some insect species and
groups have a much greater propensity to drift than
others. Mayflies, especially Baetis, and larval chironomids are perhaps the most common insect drifters.
It was once thought that when an insect entered
the drift it was essentially lost from the system. Although estimates of drift distance vary widely, most
experimental studies in small to mid-sized streams have
shown that insects drift only a short distance (a few
meters or less) during any drift event (Allan and Feifarek, 1989; Wilzbach and Cummins, 1989). However,
working in a high-order blackwater river with an important but widely dispersed snag habitat, Benke et al.
(1986) found that insects drifted between snags, a drift
distance on the scale of kilometers.
Insects may drift passively or inadvertently if they
are dislodged, such as might occur during a high discharge event or in response to an anthropogenic disturbance. However, most experimental studies have suggested that most drift is active, or a behavioral
response to some stimulus. This can occur catastrophically due to chemical spills, or on a smaller scale in response to depleted food resources (Kohler, 1985) or the
presence of a predator (Peckarsky, 1980). It was once
thought that drift was a density-dependent response,
such that drifting insects represented a portion of the
population that exceeded the carrying capacity of the
stream (Waters, 1972). However, empirical studies of
drift have yielded little support for density dependent
behavior (e.g., Hinterleitner-Anderson et al., 1992).
In most streams, drift densities are much higher at
night than during daylight hours. Because fish feed
heavily on drifting invertebrates, nighttime drift behavior
746
A. E. Hershey and G. A. Lamberti
Using 15N as a tracer, adult Baetis mayflies were
found to fly approximately 2 km upstream. The figure shows that
although adults have a lower 15N signature than nymphs, those
captured from upstream of the 15N dripper are considerably more
labeled than background. An integral mixing model showed that this
label reflected mayflies from 2 km downstream (hypothesized to be
females) mixing with mayflies from upstream (hypothesized to be
males). [From Hershey et al., 1993.]
FIGURE 12
FIGURE 11
Baetis mayflies enter drift actively, not passively, as a
result of greater activity on upper surfaces of substrates at night.
Kohler (1985) showed that only food abundance and time of day affected drift entry, and only when no food was present was there any
drift entry during the day. The figure shows the results from food
abundance experiments. [Reprinted from Kohler, 1985.]
provides invertebrates with more protection from fish
predation. Because many macroinvertebrates forage
more on the upper surfaces of rocks at night, it was once
thought that dislodgement was simply more likely at
night. However, Kohler (1985) found that well-fed Baetis
foraged only at night on tops of stones while starved
nymphs foraged during both day and night, but drifted
similarly to fed nymphs. Thus, activity during foraging
was not related to drift (i.e., they were not accidentally
dislodged) and drift entry was active (Fig. 11).
Drift clearly is an important mechanism for invertebrate dispersal, especially for early larval instars. It is also
an important means for recolonizing reaches that are impacted by natural or anthropogenic disturbance (see
Cairns, 1990). However, if there were no upstream
movement of macroinvertebrates, downstream drift
could result in a net displacement of populations, net depletion of upstream reaches, and loss of invertebrate-mediated ecosystem functions in depleted areas. The colonization cycle proposed by Müller (1954) states that
insects displaced downstream as larvae later fly upstream
as adults to oviposit, thereby repopulating depleted areas.
Upstream flight has been observed for some species (e.g.,
Madsen et al., 1973) and quantified for a Baetis population in arctic Alaska (Fig. 12; Hershey et al., 1993).
E. Saline Habitats
Insects have enjoyed only limited success in colonizing the marine environment, and only water striders
of the genus Halobates, which are surface dwelling, are
truly oceanic (Spence and Andersen, 1994). The reasons for this are unclear, but have variously been attributed to physiological stresses associated with respiration and osmoregulation, physical stress of wave
action, competition for habitat and resources with
established species, and predation (see recent reviews
by Ward, 1992; Wallace and Anderson, 1996). Because
several species of beetles, true flies, and caddisflies have
successfully colonized intertidal zones and brackish
waters, it seems unlikely that physiological stresses
alone have hindered radiation into the marine realm.
Similarly, because many insects inhabit wave-swept
lake shores and stream riffle habitats, and because
there are many low energy marine habitats, physical
stress also seems unlikely to be a major hindrance.
These considerations suggest that competitive and
predatory pressures from marine phyla, which have
been established in marine habitats much longer, are
the most likely mechanism that has precluded high insect diversity in marine habitats (Usinger, 1956; Ward,
1992; Wallace and Anderson, 1996). Alternatively,
Williams and Feltmate (1992) suggested that insects
may be in the early stages of invading marine systems,
such that they may yet become established there after
many more millennia.
18. Aquatic Insect Ecology
747
FIGURE 13
Characteristics of a stream insect community along a natural thermal gradient
emanating from a California hot spring, depicting total density, species richness (diversity),
and total biomass. [Modified from Lamberti and Resh, 1985.]
F. Specialized Habitats
Aquatic insects have invaded many specialized microhabitats. The species that comprise these communities
often have morphological, behavioral, and life-history
adaptations which are unique to those habitats and reflect the remarkable phenotypic plasticity of the Insecta.
Some of the better studied examples of these habitats include treeholes, hot springs, pitcher plants, and caves and
other subterranean environments (see Ward, 1992).
Mosquitoes are particularly well adapted to specialized ephemeral habitats such as treeholes, tires, rain
gutters, and other containers, that catch rainwater
(Merritt et al., 1992). Filter feeding is the most common mechanism for acquiring food in such habitats,
but detrital and bacterial particles may also be gleaned
from leaf and bark surfaces, and a few species are
predatory (Merritt et al., 1992). In treehole habitats, a
combination of leaf litter quality and quantity and
stemflow nutrients influence mosquito productivity
(Carpenter, 1982, 1983), but field experiments by
Walker and Merritt (1988) found no evidence that
quantity of leaf detritus was limiting.
Aquatic insects that inhabit subterranean habitats,
including cave pools and streams, underground rivers,
and extensive hyporheic zones, share some morphological characteristics. These insects, like other subterranean invertebrates, lack pigment, have reduced or absent compound eyes and ocelli, and possess reduced
wings (Ward, 1992). Diptera and Coleoptera are perhaps the best represented among subterranean insects
although representatives of other groups also have been
reported (see Ward, 1992). The trophic ecology of
these insect communities is dependent on organic matter imported from surface ecosystems. Most common
among these nutritional sources are flocculated DOM
(dissolved organic matter), entrained FPOM, bacteria
(which utilize DOM), and bat guano.
Geothermal springs are another specialized habitat
that has been exploited by some insects. Hot springs tend
to have representatives from the Diptera, Coleoptera,
and Odonata (see Ward, 1992). In addition to the high
temperatures, these habitats also have high mineral content, which may pose toxicity problems for some species.
However, Lamberti and Resh (1983a, 1985) showed that
thermal effects were more important than chemistry in
limiting insect diversity in a northern California stream
receiving natural hot spring inputs (Fig. 13).
III. AQUATIC INSECT LIFE HISTORIES
Aquatic insects show tremendous diversity in their
life-history patterns. This diversity includes variation in
reproductive and dispersal strategies, life-cycle length,
growth rate, developmental strategies, and phenology
of the various life-history stages. This great variety
serves to separate taxa seasonally and spatially, thus
providing the basis for the varied and dynamic nature
of aquatic insect community composition.
Virtually all aquatic insects have an aerial adult
phase. Copulation usually occurs outside of the aquatic
748
A. E. Hershey and G. A. Lamberti
environment, and eggs are laid on or in the water for
most species. Typically, the larval stage dominates the
life cycle; the adult stage is very short and does not always involve feeding. Notable exceptions to this rule
are the Odonata (dragonflies and damselflies), that are
long-lived and voracious predators as adults. Dragonflies and damselflies also have long flight periods, feeding on insect swarms and using this energy for flight,
food capture, and reproduction. Swarms of dragonflies
often are thought to be important for control of mosquitoes and blackflies. Many dipterans also feed as
adults. Mosquitoes and blackflies are notorius pests,
but are also vectors of diseases such as malaria, encephalites, dengue, Rift Valley Fever, river blindness,
and many others that infect and kill millions of people
worldwide (e. g., Renshaw et al., 1996; Jacobs-Lorena
and Lemos, 1995; Kidwell and Wattam, 1998). With
global warming, the geographic distribution of these
diseases is expected to increase (Brown, 1996).
For many aquatic insects, the adult phase is often
important to dispersal between lentic habitats, and permits upstream oviposition (see Müller, 1954; Hershey
et al., 1993), and oviposition between watersheds (Wallace et al., 1986) in lotic habitats. Many aquatic beetles
and bugs have extended adult life spans and are aquatic
in both larval and adult stages of the life cycle, although
dispersal still is accomplished by the adult stage. Dispersal varies tremendously among insects and has been
studied thoroughly for only a few taxa. For example,
Baetis mayflies in a tundra river in arctic Alaska have
been shown to fly approximately 2 km upstream to lay
eggs (Hershey et al., 1993). Even larval insects can
move substantial distances, aside from drift. For example, late-instar larvae of the grazing caddisfly Dicosmoecus gilvipes can crawl up to 25 m per day in search of
algal food (Hart and Resh, 1980). Nymphs of the lotic
mayfly, Leptophlebia cupida, move upstream at an
average rate of 10 m “per hour” during seasonal migrations between habitats (Hayden and Clifford, 1974).
The frequency with which an organism completes
its life cycle is referred to as “voltinism.” Depending on
environmental constraints, especially temperature and
nutrition, as well as evolutionary factors, aquatic insects
may be univoltine (one generation per year), multivoltine (more than one generation per year; e.g., bivoltine
two per year, trivoltine three per year), semivoltine
(2-year life cycle), or merovoltine (3- or more-year life
cycle). Generally, larger insects will have longer life cycles than smaller species. For example, in the Kuparuk
River in arctic Alaska, the relatively small Baetis
mayflies are univoltine, but the larger Brachycentrus
caddisflies are merovoltine (Hershey et al., 1997). Univoltine development is by far the most common, especially in temperate regions (see Wallace and Anderson,
1996). However, even in arctic streams, where the
stream bed is frozen for almost nine months, most
species are univoltine (Hershey et al., 1997). Arctic lakes
have chironomid populations with life cycles ranging
from 1 to 4 years, but univoltine development predominates (Hershey, 1985a). One arctic pond species of chironomid has a 7-year life cycle (Butler, 1982). Temperate
regions display similar variation. The wood-boring
aquatic beetle Lara avara has about a 5-year life cycle,
likely due to the low nutritional value of wood (Anderson and Sedell, 1979). At the other extreme, some herbivorous chironomid midges in warm desert streams can
complete their life cycle in as little as two weeks, a distinct advantage in desert streams where floods scour organisms and resources (Fisher and Gray, 1983).
Variation in growth rate is also important in distinguishing insect species, separating cohorts, and providing structure to aquatic insect communities. Within
physiological constraints, temperature and food are the
major factors affecting growth for most taxa. These
factors vary widely within, and between, lentic and
lotic ecosystems, as well as geographically.
In cool climates many aquatic insects have life cycles
with synchronized development (Fig. 14). Environmental
factors, especially temperature and photoperiod, as well
as evolutionary factors, serve as proximal cues that synchronize life cycles (Butler, 1984). For example, many insects will overwinter in the egg stage, and hatch in the
spring. Others may overwinter as larvae, emerging in the
spring when the appropriate temperature and / or photoperiod cues are present. Although there may be several
evolutionary as well as environmental mechanisms contributing to synchronous development, and contributing
factors vary among species (see Butler, 1984), the result is
that within a species, individuals in a “cohort,” or age
class, will emerge during a very short interval, enhancing
the probability of mating success. Synchrony occurs
through two general paths. Larval hatching and growth
may be timed such that all individuals within a cohort
are of identical or nearly identical size throughout their
development and emerge together. This is common in
many caddisflies, such as the widely distributed
Helicopsyche borealis (Resh et al., 1984) and the large
limnephilid Dicosmoecus gilvipes (Lamberti and Resh,
1979). Alternatively, hatching and larval development
may not be synchronous but instar-specific growth rates
still may result in synchronized emergence (Lutz, 1974).
Within a community, co-occurring species typically
exhibit a variety of patterns of voltinism and synchrony. For example, 14 species of dragonflies in a
temperate pond were found to have several different
developmental patterns (Wissinger, 1988). Epitheca
cynosura was univotine with synchronous development
(Fig. 15A), while one of its congeners, E. princeps, also
18. Aquatic Insect Ecology
FIGURE 14 Examples of life cycle synchrony of stream macroinvertebrates in response to temperature and photoperiod cues. In
temperate North America, many species overwinter as larvae (left).
Larval growth and maturation occurs at approximately the same rate
for the entire cohort, such that emergence will also be synchronized.
Other species overwinter in the egg stage, then begin larval development in the spring (right). These developmental patterns increase the
probability that adults will find mates, thus enhancing reproductive
success. Larval instars indicated as: F, final instar; F-1, penultimate
instar; etc; and P, pupae. [From Anderson and Bourne, 1975.]
749
FIGURE 15 Developmental patterns of Epitheca spp and Libellula
lydia illustrate that even closely related species from the same habitat
can exhibit divergent life-history patterns. (A) Univoltine synchronized development. (B) Semivoltine synchronized development. (C)
Semivoltine asynchronous development. [From Wissinger, 1988.]
750
A. E. Hershey and G. A. Lamberti
FIGURE 16 The two major forms of insect development seen in aquatic insects. In gradual metamorphosis (left), or hemimetabolous development, immature forms are insectlike. Adult characteristics develop gradually through the larval instars, and wings inflate in the final molt to the adult form.
Insects demonstrating complete metamorphosis (right), or holometabolous development, have several
larval instars that are less insectlike. Adult characteristics develop as a result of radical restructuring
of the anatomy and physiology during the pupal instar. [From McCafferty, 1981.]
was synchronized but was semivoltine (Fig. 15B). Yet
another congener, E. simplicollis, was sometimes bivoltine. Asynchronous univoltine development was exhibited by Libellula lydia (Fig. 15C), while other species
showed different combinations of synchrony, asynchrony, and mixed voltinism (Wissinger, 1988).
Developmentally, most insects can be categorized
as either “hemimetabolous” or “holometabolous” (Fig.
16). Collembola, considered “ametabolous,” have a developmental pattern that is distinctive from other insects and is phylogenetically primitive. This has led
some entomologists to exclude it from the Insecta
(Borror et al., 1989). The insect orders Ephemeroptera,
Odonata, Heteroptera, and Plecoptera, are hemimetabolous. The aquatic immature forms, with few exceptions, respire using gills, and although they may differ
considerably from adults, they are clearly insectlike
(Fig. 16). The immature stages are referred to as
nymphs or larvae. A newly hatched larva is referred to
as a first instar, and larval development proceeds
through a series of instars. The number of instars varies
widely among families, but variability also may occur
at the species level depending on environmental conditions (Borror et al., 1989). Most aquatic insects are
holometabolous, which means they undergo complete
metamorphosis (Fig. 16). The immature instars are referred to as larvae and do not resemble the adults. Larval appearance is variable, but is usually not insectlike.
Commonly larvae are described as wormlike, grublike,
or caterpillarlike. A pupal stage that follows the last
larval instar is characterized by development of the
adult form in a protected habitat or cocoon. The insect
does not feed during this stage, and appears to be quiescent. However, metabolic activity is very high as
18. Aquatic Insect Ecology
larval tissues are reorganized to form the morphologically and physiologically different adult form.
IV. INSECT-MEDIATED PROCESSES
Insects are important consumers and important
prey items in virtually all aquatic habitats, thus are intimately involved in the flows of matter and energy that
occur in these ecosystems. Insects also exert top – down
controls on nutrient cycles, primary productivity, decomposition, and translocation of materials, and affect
relative abundances of other consumers in lentic as well
as lotic ecosystems. Specific relationships between insects and aquatic resources that are of particular interest include detritivory, grazing, and predation.
Processes that integrate one or more of these specific
relationships include secondary production and stream
drift. All of these processes are shaped by physical
habitat and life-history constraints.
A. Detritivory
Detritivory, the feeding on decaying organic matter,
is a major insect-mediated process in both lentic and
lotic systems, although it has received more attention in
lotic studies. The relative importance of allochthonous
versus autochthonous detritus in aquatic ecosystems is
widely variable, and size of the water body is the major
determining factor. Consumer resources in small lentic
systems and low-order streams are typically dominated
by allochthonous inputs, whereas lakes and middleorder streams (see Vannote et al., 1980) are usually
dominated by autochthonous production. When detritus is plentiful, however, detritivory by insects is usually important to ecosystem function.
Detritus forms a major habitat for aquatic insects in
ponds and many wetlands. The coarseness of the
detrital substrate provides habitat for large aquatic insects, such as odonate nymphs, which prey on small
detritivorous chironomids, mayflies, and microcrustaceans. For some large insects, detritus serves as a food
source, although availability of detritus seldom limits
wetland insect abundance or production, and microbial
decomposition appears to be more important than insect-mediated decomposition in many wetlands (Batzer
and Wissinger, 1996). Detritus is divided into two major
fractions: coarse particulate organic matter (CPOM —
that portion 1 mm in size) and fine particulate organic
matter (FPOM — the portion that is 1 mm).
Low-order streams are strongly influenced by the
input of terrestrial organic matter from adjoining
ecosystems. In temperate streams, much of this detritus
enters as a seasonal (autumnal) pulse of leaves, which
751
is then processed in the stream by microbial and
macroinvertebrate communities, and serves as the major energy source for many stream consumers (Fig. 9).
Life cycles of many detritivorous insects are cued to
this seasonal pulse of leaves such that they develop
through the fall and winter when their food resource
also is plentiful. Microbes, which colonize the detritus,
form an integral part of the CPOM complex and are
nutritionally important to shredders (Anderson and
Sedell, 1979; Cummins and Klug, 1979). Dominant
shredder taxa vary according to stream type and biogeography. Shredders are found in many of the aquatic
insect orders (Table I). Some of the more conspicuous
insect shredders are pteronarcid and capniid stoneflies,
larval craneflies, and several families of caddisflies.
Low-order streams often have 30 – 50% shredders in
their macroinvertebrate communities (Fig. 10; see also
Fig. 6 in Chapter 2), as was found in Oregon streams
(Hawkins and Sedell, 1981), by virtue of their closed
canopy and high input of CPOM per unit of stream
surface (discussed in Vannote et al., 1980).
In a first-order mountain stream, Wallace et al.
(1997) used an ecosystem-level experiment (litter exclusion by suspending a net over a small stream) to
demonstrate that terrestrial litter supported the vast
majority of insect secondary production in the stream.
Interestingly, insects and other consumers occupying a
moss microhabitat in the stream did not respond to
litter exclusion, suggesting that moss inhabitants comprised a food web that was independent of that supported by terrestrial detritus (Wallace et al., 1997). In a
different study, experimental reduction of shredders in
a stream using an insecticide resulted in lower CPOM
decomposition (20 – 40% that of a reference stream)
and 5- to 15-fold lower FPOM transport (Wallace
et al., 1982). Clearly, insects are of critical importance
to CPOM processing in streams.
Formation of FPOM from CPOM by shredders during their feeding is an important process. FPOM is also
formed by several other mechanisms, and is dynamic in
stream food webs. All consumers produce feces, which
are an important component of FPOM (Shepard and
Minshall, 1981; Hershey et al., 1996). Bacteria use
DOM, thereby incorporating it into bacterial biomass,
which itself constitutes a portion of the FPOM pool
(e.g., Meyer, 1990). Bacteria produce extracellular materials, which are sloughed into the FPOM pool (e.g.,
Wotton, 1988). FPOM is produced by flocculation of
DOM due to a variety of physical and chemical
processes (Dahm, 1981; Petersen, 1986; Wotton, 1988;
Ward et al., 1994). The mass of FPOM usually exceeds
that of nonwoody CPOM by about an order of magnitude (Wallace and Grubaugh, 1996) and turns over
much more rapidly. Insect collector-filterers and
752
A. E. Hershey and G. A. Lamberti
collector-gatherers that use this resource are important
components of the macroinvertebrate community in all
streams (Fig. 9; Vannote et al., 1980; Hawkins and
Sedell, 1981). However, because FPOM is formed by
many processes, it varies widely in quality; some FPOM
is highly labile, but much is refractory (Newbold et al.,
1983). Its retention in streams is also high, but varies
considerably due to physical and / or chemical characteristics of the particles (Miller et al., 1998). The variety of
collectors that use FPOM also is very high (Table I).
DOM is generally considered to be the exclusive
resource of stream microbes, which assimilate labile
components of DOM into microbial biomass (see Lock
et al., 1984; Meyer, 1990). However, larval blackflies
also ingest DOM (Fig. 17, and see Wotton, 1976) and
FIGURE 17
Ingestion of DOM by larval blackflies. Heads and bodies of black flies which had been permitted to feed for 10- or 60-min
on 0.2-m filtered 14C-DOM were assayed separately for incorporation of labeled material. After 10 min, similar activity was seen on
bodies of live and dead larvae, but higher activity was seen on heads
of live larvae. This indicated that activity of live larvae enhanced
accumulation of 14C from DOM onto the head. After 60 min, considerably more material was present in the body, indicating that labeled
DOM was being packed into the gut during feeding. Each set of bars
shows results from a separate chamber (two live chambers and one
dead chamber). [From Hershey et al., 1996.]
this process can be important to the size and quality of
the FPOM pool downstream of blackfly aggregations
(Hershey et al., 1996). DOM ingestion by blackflies includes some labile material, such as bacterial exopolymer (Couch et al., 1996), and thus such feeding may be
a nontrivial source of nutrition for these aquatic insects. Although direct DOM ingestion by stream insects
is not thoroughly understood, its importance is likely
limited to a few taxonomic groups.
Large woody debris (LWD) often forms an important pool of detritus in stream ecosystems (Anderson
and Sedell, 1979; Bilby and Ward, 1989; Swanson et al.,
1982; Van Sickle and Gregory, 1990). Wood may cover
over 25% of the beds of small streams in old-growth
forests (Anderson and Sedell, 1979). However, only
very specialized insects use this material as food, and
these likely do not ingest quantities that are significant
to wood decomposition (e.g., Wallace and Anderson,
1996). LWD is important as macroinvertebrate habitat.
Benke et al. (1984) showed that much of the secondary
production in a southeastern blackwater stream was associated with woody snags. In addition, the insect community on the snags was distinct from other substrates.
LWD also serves as an important mechanism for retaining CPOM and FPOM resources that are used by insects (Benke et al., 1984). In addition, LWD is a major
factor defining pool and riffle habitat within a stream,
which determines many aspects of macroinvertebrate
community structure. Wood itself has low nutritional
value, but the microorganisms that coat submerged
wood (e.g., algae, bacteria, fungi, protozoans) provide a
rich food resource for macroinvertebrates.
A few invertebrates (“wood gougers,” a special
group of shredders) are adapted to feed directly on the
wood and include the cranefly larva Lipsothrix and the
elmid beetle Lara avara (Anderson and Sedell, 1979;
Wallace and Anderson, 1996). These taxa bore into
and feed strictly on wood and have very long life cycles
(3 – 6 years) as a result of the low nutritive value of
their food. Examining a piece of water-logged wood
will reveal the tunnels of these borers and often the insects themselves. Wood is also used by some larval caddisflies for case construction. Heteroplecton californicum, a caddisfly that can be found in shallow pools,
hollow out twigs to construct portable cases (Fig. 18).
When the larvae move, their cases appear to be moving
twigs, and the larvae themselves are not visible.
B. Grazing
Benthic primary production is a major process in
virtually all shallow aquatic ecosystems (Lamberti,
1996). As a consequence, grazing insects functioning as
primary consumers can be found in nearly all of those
18. Aquatic Insect Ecology
753
FIGURE 18
Wood cases of the caddisfly Heteroplectron californicum next to shredded alder
leaves. Note opening chewed by caddisflies at upper end of twigs; larva resides inside the twig
within a silk-lined tunnel. [Photograph by Jon Speir, reproduced from Anderson, 1976a.]
ecosystems except, perhaps, where extreme physical
conditions predominate (Lamberti and Moore, 1984).
Aquatic primary producers include algae, rooted
macrophytes, and bryophytes (mosses and liverworts).
Benthic algae frequently are the dominant plants in
streams and along rocky, wave-swept littoral zones,
whereas rooted macrophytes become more important
in the finer sediments of lakes and large rivers where
they are exploited for both habitat and food by insects.
Fine sediments may also be covered with a layer of algal “epipelon,” which is readily grazed by insects.
Mosses can be locally important, especially in streams,
and because of their fibrous texture and long-lived nature they provide excellent habitat for a variety of insects (Glime and Clemons, 1972; Suren and Winterbourn, 1992). Where mosses line the rock walls and
spray zones of waterfalls, often a specialized insect
fauna develops termed “torrenticolous” insects. In the
discussion below, we will mostly concern ourselves
with grazers of periphyton and macrophytes.
Scrapers can be found in six orders of aquatic insects (Table I), and for several of those orders
(Ephemeroptera, Trichoptera, and Lepidoptera) algal
consumption is a major way of life. Grazers often possess elaborate mechanisms to remove periphyton (attached algae, microbes, and associated organic matter)
from surfaces (Lamberti et al., 1987a). Many species of
mayflies bear stout bristles on the labial or maxillary
palps that are effective in removing periphyton from
cracks and crevices. These bristles increase in length
with successive instars, thereby allowing older larvae to
remove larger particles while younger larvae remove
smaller particles, effectively partitioning the resource
by age. Many caddisfly larvae have robust or scoopshaped mandibles for removing or “spooning” periphyton into their mouths. Some limnephilid caddisflies,
such as Dicosmoecus, also use their tarsal claws to
gather algae into small piles for ingestion (Lamberti
et al., 1987a). Orthocladius chironomids have stout
mandibles lined with 4 – 5 teeth that wear down over
time from scraping (Saponis, 1977); fortunately, molting periodically provides them with a new set of teeth.
Even presumed shredders, such as cranefly larvae (Tipulidae), possess setae on the cutting surfaces of their
mandibles that may help to collect algae from leaf surfaces, thereby allowing these insects to function as a
facultative grazers (Pritchard, 1983). For shredders, the
microorganisms, including algae, that grow on decaying leaves may be the most nutritional component of
the leaves (Cummins et al., 1989).
Grazing has been the focus of much study in streams (reviewed by Steinman, 1996, and Feminella and Hawkins,
1995, for field studies and by Lamberti, 1993, for laboratory
stream studies) and to a lesser extent in lakes (reviewed by
Lodge, 1991; Newman, 1991). In streams, benthic algae will
grow on virtually any substrate exposed to light, including on
benthic insects themselves or their cases (e.g., Pringle, 1985;
Hershey et al., 1988; Bergey and Resh, 1994). Many streams,
however, have a low standing crop of benthic algae, sometimes barely perceptible to the naked eye. In comparison with
obvious pools of dead organic matter (e.g., leaves and wood),
it is tempting to dismiss the significance of algae and thereby
754
A. E. Hershey and G. A. Lamberti
grazing. However, a specific standing crop of algae can support a biomass of herbivores 10–20 greater than its own
because of high algal turnover rates (McIntire, 1973), sometimes referred to as an “inverted trophic pyramid.” For example, Mayer and Likens (1987) found that an abundant caddisfly (Neophylax aniqua) consumed mostly periphyton in a
small, heavily shaded New Hampshire stream whose energy
base was previously thought to be almost totally detrital
(Fisher and Likens, 1973). Still, periphyton is often limiting to
the growth of insect grazers, and competition for algal
resources has been demonstrated (e.g., McAuliffe, 1984;
Lamberti et al., 1987b).
Among the stream insects, caddisflies (Trichoptera)
and mayflies (Ephemeroptera) tend to be highly conspicuous grazers although small chironomid midges
(Diptera) may be equally important herbivores due to
their ubiquity, high densities, and short generation
times (Berg and Hellenthal, 1992). Insect grazers have
the ability as individual species or groups of taxa to exert large effects on the abundance, productivity, and
community structure of benthic algae. Often, a single
species can exert primary control over periphyton. For
example, Helicopsyche borealis caddisflies consumed
95% of the algal standing crop in a northern California stream (Lamberti and Resh, 1983b). Experimental
exclusion of these caddisflies produced a mat of filamentous algae almost 30-mm thick (Fig. 19). In another study, the large caddisfly Dicosmoecus gilvipes
consumed about 60% of benthic algal production, and
FIGURE 19
most of the remaining algae were dislodged and
washed away by the grazing activities of these
“sloppy” feeders (Lamberti et al., 1987a). Furthermore, these large grazers can have negative effects on
other benthic invertebrates, which they “bulldoze” and
thereby displace as they harvest periphyton (Hawkins
and Furnish, 1987; Lamberti et al., 1992).
Tightly evolved associations between plants and
herbivores are well documented in terrestrial ecosystems (Crawley, 1983). In aquatic ecosystems, however,
there are few examples of host-specific associations between aquatic insects and benthic algae, for reasons
that are still unclear (Gregory, 1983). One possible example is found in North American streams, where a
mutualistic relationship may exist between the chironomid midge Cricotopus nostocicola and the blue-green
alga Nostoc parmelioides (Brock, 1960). The first-instar midge larva finds and enters a small, globular
colony of Nostoc and begins feeding on Nostoc cells, in
the process changing the colony morphology to an earlike form about 1 cm in diameter (Fig. 20). These darkgreen colonies often are quite obvious on rocks, and
the single larva can be seen clearly within the colony.
The midge grows and pupates within the colony, finally
emerging as an adult. The association also may benefit
the colony, which has a higher photosynthetic rate
when the fly is present (Ward et al., 1985; Dodds,
1989). Amazingly, the midge will reattach the colony
if it becomes dislodged from the substrate during a
Effects of grazing by the caddisfly Helicopsyche borealis on algae in Big
Sulphur Creek, California. Elevated but submersed platform in the foreground was used
to exclude benthic caddisfly larvae; tiles in midground and rocks in background were
freely grazed by caddisflies at densities 1000 / m2. Direction of water flow is from right
to left. (A) Beginning of experiment. (B) After 5 weeks elapsed. Note dense growth of filamentous algae on ungrazed platform tiles. [From Lamberti and Resh, 1983b.]
18. Aquatic Insect Ecology
FIGURE 20
Colonies of the blue-green alga Nostoc parmelioides containing the midge larva Cricotopus
nostocicola. Upper panel: Nostoc “ear” form with midge larva in left side of colony; Lower panel: Nostoc
colonies attached to a rock. Bar 5 mm. [Photographs by Louis Nelson, reproduced from Ward et al., 1985.]
755
756
A. E. Hershey and G. A. Lamberti
disturbance (Brock, 1960; Dodds and Marra, 1989),
which doubtless benefits both the colony and the larva.
Another example, although probably not mutualistic, is
the association between the microcaddis fly Dibusa angata and the freshwater red alga Lemanea australis
(Resh and Houp, 1986). Larval instars 1 – 4 of Dibusa
are found among the basal holdfasts of Lemanea,
where they consume epiphytic diatoms. Fifth-instar larvae construct cases made of Lemanea and consume
only Lemanea tissue. There is no known benefit of this
association to the alga.
Many insects consume living macrophyte tissue, a
process that generally has been understudied and underappreciated (see review by Lodge, 1991). Unlike for
algae, there appear to be a number of close associations
between insects and aquatic macrophytes, perhaps because the large size of rooted plants makes specialization more feasible than for microscopic algae. The major insect orders involved in macrophyte grazing are
the Trichoptera, Lepidoptera, Coleoptera, and Diptera.
It is perhaps not surprising that rooted hydrophytes are
eaten by the more advanced insect orders, as their terrestrial insect relatives are also mainly plant eaters. Indeed, it has been speculated that over evolutionary time
terrestrial insects followed their host plants as they
gradually invaded the water to eventually become true
hydrophytes. Furthermore, parasitoids of those insects
may have followed their hosts, as evidenced by the hymenopterans that now parasitize aquatic lepidopterans.
Insect grazers can consume up to 100% of the standing
stock of macrophytes, although the damage is usually
much less than that (Lodge, 1991). It is more common
to see the grazing scars of insects, such as beetles and
moths, on the surfaces of floating leaves and other
plant structures. Lepidopterans are known to feed on
aquatic plants in numerous ways, including leaf mining, stem or root boring, foliage feeding, and flower or
seed consumption (Lange, 1996). Coleopterans likewise exhibit a wide variety of feeding modes; leaf beetles (Chrysomelidae) are particularly common herbivores of water lillies, where their larval galleries are
obvious on the floating leaves (Cronin et al., 1998).
Even though beetles may not consume large amounts
of leaf tissue, their feeding activity can increase nutrient
uptake and primary production by the plant and accelerate decoposition (Wallace and O’Hop, 1985).
C. Predator – Prey Interactions
Many aquatic insects function as predators, and
predators are found in nearly all of the major groups of
aquatic insects. A few groups, notably the Odonata,
aquatic Heteroptera, and Megaloptera, are strictly
predators, whereas other groups are more heteroge-
neous in their trophic affiliations, but may have important subdivisions that are predators. For example,
among the Plecoptera, most members of the families
Perlidae, Perlodidae, and Chloroperlidae are primarily
predators (Stewart and Harper, 1996). Within the Trichoptera, members of the family Rhyacophilidae are
free-roaming predators, which may inflict particular
damage to larval black fly populations (Wheeler, 1994).
Many of the sedentary, net-spinning Hydropsychidae
are facultative predators (Benke and Wallace, 1980),
and some larval Limnephilidae become opportunistic
predators in later instars (Anderson, 1976a). Many
Coleoptera and Heteroptera function as aquatic predators in both the immature and adult forms.
The role of predators in controlling the distribution and abundance of prey is a central question in
aquatic ecology, and their role in determining aquatic
insect communities also has received considerable attention. Predator and prey taxa are subjected to different physical constraints in various lentic and lotic
ecosystems (see above), and these physical constraints
interact with the predation process to produce different
community types. Because predator – prey interactions
do not always occur at the same spatial and temporal
scales as abiotic factors that affect distribution and
abundance of organisms, it is sometimes difficult to extrapolate predator – prey observations to community
composition (see Peckarsky et al., 1997).
Predators acquire prey using different behavioral
strategies (see Peckarsky, 1982). Many predators, such
as stoneflies, actively search for their prey (“hunters”).
Ambush or “sit-and-wait” predators wait until a prey
item is within range before striking. Odonate larvae often use this strategy. Once the prey is captured predators typically either engulf their prey intact, in bites, or
may feed only on the internal fluids. Fluid feeders use
specialized piercing (haustellate) mouthparts for injecting enzymes into their prey and sucking out the partially digested body. Clearly the relative sizes of predators and prey, as well as the mode of feeding used by
the predator, are important mechanical constraints governing predator – prey interactions. Engulfing predators
are limited to prey small enough to be subdued and
swallowed intact (see Hershey, 1987), whereas piercing
predators have access to a broader variety of prey sizes
that are large enough to be handled and to accommodate the piercing mouthparts.
In a synthesis of predator effects in lentic communities, Wellborn et al. (1996) characterized control of
community structure as falling along a gradient of physical and biotic control, with two major transitions (Fig.
21). Temporary ponds and lakes provide two extremes
along a habitat size and permanence gradient, and two
major transition thresholds occur along this perma-
18. Aquatic Insect Ecology
757
FIGURE 21
Aquatic communities are controlled by both biotic
and abiotic factors. Two major transitions involving predator control
of community structure are conceptualized to occur (Wellborn et al.,
1996). A permanence transition reflects when the wet phase of the
hydroperiod is sufficiently long to support longer-lived predatory insects. A predator transition occurs when habitats contain fish, which
then eliminate many predatory insects. [From Wellborn et al., 1996.]
nence gradient, which provide the template for interaction of physical factors and predator control of insect
communities (Fig. 21; Wellborn et al., 1996). A permanence transition (Fig. 21) along the gradient controls
predator presence or absence. In highly ephemeral habitats, most predatory insects are excluded (Batzer and
Wissinger, 1996; Wellborn et al., 1996, and see above).
As a result, predator – prey interactions are relatively
unimportant and the insect community is controlled by
physical factors (Wellborn et al., 1996), primarily
drought cycles (Hershey et al., 1998, 1999). Once the
permanence transition is crossed, predatory insects become abundant and control various aspects of prey
community composition (e.g., Batzer and Resh, 1996).
These habitats tend to have the highest diversity of insect species (see Batzer and Wissinger, 1996). A predator transition occurs when habitats are large enough
and deep enough to support fish (Fig. 21; Wellborn
et al., 1996). Fish are size-selective predators, typically
eliminating large predatory taxa, which have a major
impact on prey diversity and community structure. For
example, in an experimental manipulation of bluegill
and pumkinseed sunfish, mean invertebrate size decreased as fish density increased (Mittlebach, 1988). But
fish may control insect prey density without dramatically affecting community composition (Bohanan and
Johnson, 1983). Fish and predatory insects interact to
increase the effect on prey compared to when either is
present alone (Johnson et al., 1996). But fish and large
predatory insects such as dragonflies have different
strategies for capturing prey (i.e., pursuit versus ambush). These alternative strategies provide contrasting
FIGURE 22 A fish exclusion experiment conducted in dense
macrophytes of a cooling water reservoir in South Carolina did not
show significant reduction of macroinvertebrates in the areas where
no fish were present. NP, no predatory fish; and P, predatory fish
present. [Reprinted from Thorp and Bergey, 1981.]
selective environments for prey; McPeek et al. (1996)
have shown that in lakes where dragonfly predation is
important, Enallagma damselfies have larger caudal
lamellae which enhance swimming speed, whereas in
fish lakes, Enallagma caudal lamellae are smaller, associated with a cryptic strategy for predator avoidance.
Habitat heterogeneity moderates the effects of fish
predation on insect communities (Gilinsky, 1984;
Hershey, 1985); and in very dense macrophytes, fish
are not important in controlling insect abundance or
species composition (Fig. 22; Thorp and Bergey, 1981;
Crowder and Cooper, 1982). Thorp (1986) concluded
that although fish native to a system may control density or biomass of benthic insects, they have little impact on such community structure characteristics as diversity or species composition. However, historical
presence or absence of fish within a community has a
major impact on community structure (Fig. 21; Thorp,
1986; Wellborn et al., 1996).
Similar to lentic studies of predation, predator –
prey interactions involving stream insects also have
been studied from two major perspectives: (1) the
potential for their communities to be structured by
fish predation; and (2) as predators interacting with,
and possibly controlling, other macroinvertebrates.
As in lentic systems, experimental studies of fish predation have not always led to the same conclusions.
In an experimental removal of trout from a rocky
mountain stream, Allan (1982) found no difference in
macroinvertebrate density or diversity where fish were
758
A. E. Hershey and G. A. Lamberti
FIGURE 23
Simplified food web found in the Eel River, California
(Power, 1990) where cascading effects have been demonstrated.
Large fish control small fish that feed preferentially on predatory
insects. This releases tube-building chironomids from control by insects, and dramatically affects the appearance of the algal mat.
[Reprinted from Power, 1990.]
removed. Similarly, Reice (1983) found no evidence of
trout control of any aspect of the stream insect communities that he studied. However, working in California’s
Eel River, Power (1990) has shown that predatory fish
control predatory invertebrates (especially damselflies)
which, in turn, control the abundance of larval chironomids, affecting both algal biomass and the physical appearance of the algal mat in the river (Fig. 23). Her
study is an example of food web control via cascading
trophic interactions (sensu Carpenter et al., 1985),
which have been well studied for pelagic systems. Although it is not clear that trophic cascades are common
in stream ecosystems, it is clear that fish predation does
not control insect abundance in all streams.
Regardless of their direct effects on prey density or
community structure, fish have important indirect effects on prey in lotic ecosystems (e.g., Wooster, 1994;
Wallace and Webster, 1996). Because of the threat of
fish predation, many insects feed more actively on rock
surfaces at night and are more prone to drift at night in
search of better food patches (Fig. 11; Kohler, 1985). It
appears that stream fishes are usually limited by their
macroinvertebrate (especially insect) prey resources.
This is suggested by Allen’s paradox (Allen, 1951), but
the strongest evidence for this comes from experimental fertilization studies that result in increased fish production (e.g., Stockner and Shortreed, 1978; Deegan
and Peterson, 1992; Richardson, 1993).
The role of predatory macroinvertebrates in
streams has been studied using a combination of gut
analyses, laboratory stream microcosms, and in-stream
experiments. Numerous studies of gut contents have
yielded a wealth of information on which invertebrate
species function as predators and which species or
groups serve as their major prey. However, insight into
their role in structuring stream communities has come
primarily from experimental studies. Although notable
exceptions exist (e.g., Power, 1990): invertebrate predators probably do not control the abundances of their
prey in most stream ecosystems because stream communities are very open to immigration and emigration
of both predators and prey (see Cooper et al., 1990).
However, indirect effects appear to be of considerable
consequence. Behavioral avoidance of predators by
prey may affect prey spatial distribution within the
substrate. For example, predatory stoneflies affect
mayfly distributions because mayflies will drift to avoid
contact with them (Peckarsky and Dodson, 1980).
When contact occurs, Baetis may further minimize the
threat of predation by projecting their cerci forward in
a “scorpionlike” posture that renders them more difficult for the stonefly to handle (Fig. 24; Peckarsky,
1980). Prey growth rates and / or fecundity can also be
altered by the presence of predators, even if survivorship is not altered (Peckarsky et al., 1993; Scrimgeour
and Culp, 1994), and shredding activity can be reduced
(Malmqvist, 1993). Alteration of growth rates may or
may not have implications for prey demographics, depending on whether adult fecundity is a function of
body size, as it is in Baetis mayflies but not in Enallagma damselflies (McPeek and Peckarsky, 1998). Prey
also respond to predators through morphological
adaptation. Ephemerella are armored against stonefly
predators (Peckarsky, 1996). In stream habitats where
Hydra are very abundant, long hairs on the midge
larva Cricotopus sylvestris greatly decreases predation
risk relative to short haired species, such as C.
bicinctus, as well as to individuals of C. sylvestris
whose hairs have been experimentally shortened (Fig.
25; Hershey and Dodson, 1987). A conceptual model
by Peckarsky (1996) suggests that resource acquisition
modes may account for investment in morphological
versus behavioral strategies for predator avoidance.
These examples show that predator – prey interactions
involving insects are diverse, and variously may alter
prey abundances, distributions, and behavior, and may
have been important in selecting for morphological
adaptations for predator defense.
D. Insects as Conduits for Energy Flow
in Aquatic Food Webs
Insects provide a critical linkage in energy flow
from microbial to vertebrate populations in aquatic
ecosystems. Food web analyses have been used to
understand these linkages, and thereby integrate organic-matter processing with community interactions
(e.g., Closs and Lake, 1994; Benke and Wallace, 1997).
The goals of food web studies for a particular system
are to identify organic matter sources for the various
18. Aquatic Insect Ecology
759
FIGURE 25
Long hairs provide C. sylvestris with protection
against Hydra, compared to short haired C. bicinctus or C. sylvestris
with experimentally shortened hairs. Hydra is a freshwater cnidarian
that preys on chironomids and other aquatic invertebrates. [From
Hershey and Dodson, 1987.]
FIGURE 24
Mayflies exhibit a defensive “scorpion” posture when
attacked by predatory stoneflies. [From Peckarsky, 1980.]
consumers and to elucidate the trophic structure of the
web. In most aquatic ecosystems, either three or four
trophic levels are present including: (1) primary producers and detritus; (2) primary consumers, including
detritivores (shredders and collectors) and grazers
(scrapers); (3) secondary consumers (predators); and
(4) tertiary consumers (vertebrate predators that consume invertebrate predators). These designations are
oversimplifications because omnivory is common, such
that many consumers occupy more than one trophic
level (see Lamberti, 1996). In stream ecosystems, the
relative importance of the detritivore food chain compared to the grazer food chain is a matter of some debate (Minshall, 1978), but this varies as a function of
stream order and riparian conditions (Hawkins and
Sedell, 1981; Cummins et al., 1989; Gregory et al.,
1991). The food web in any particular aquatic ecosystem will reflect the combination of factors altering resource and invertebrate abundance and distribution.
Thus, wide variation in food web structure occurs. Gut
analyses are often used to delineate trophic structure
(e.g., Hershey and Peterson, 1996). However, some
food items are difficult or impossible to recognize,
while others may be overlooked due to rarity or
760
A. E. Hershey and G. A. Lamberti
temporal variability. The gut analysis approach is impractical for studies involving fluid-feeding predators
because sclerotized parts are not ingested, thus prey
identity cannot be determined.
Recently, stable isotopes have been used to study
food-web relationships in both lotic and lentic systems
(e. g., Fry and Sherr, 1984; Minagawa and Wada, 1984;
Peterson and Fry, 1987; Fry, 1991; Peterson et al.,
1993; Junger and Planas, 1994; Schuldt and Hershey,
1995; Whitledge and Rabeni, 1997; Vander Zanden
and Rasmussen, 1999). One useful aspect of carbon stable isotopes is that the various organic matter sources
for consumers often have different relative abundances
of the two stable isotopes of carbon, 13C and 12C. Because the relative abundances of these isotopes change
only slightly as the organic matter is processed by various consumers, the difference between the ratio 13C to
12
C in a consumer and a standard (or value, expressed
as parts per thousand, or per mil) can be used to infer
the consumer’s food source. Recently, 15N has been used
extensively in enrichment studies of lotic and lentic
ecosystems to serve as a tracer of consumer food
sources as well as nitrogen-distribution through the system (Kline et al., 1990; Hershey et al., 1993; Wolheim
et al., 1999). 15N also is used in trophic studies. 15N and
14
N change as food is processed by consumers because
the various metabolic processes use, or fractionate,
these isotopes differently. The net result is that with
each trophic level, consumers become enriched (by
about 3 per mil) in 15N relative to 14N, as 15N is retained in the tissues and 14N is excreted. Thus, consumers in a food web can generally be assigned to a
trophic level even if their precise food resource is not
known (see Fry, 1991; Peterson et al., 1993). However,
the del 15N value of the base of the web may vary
between systems, thus should be measured for each
system where del 15N is being studied in consumers
(Vander Zanden and Rasmussen, 1999).
When organic matter sources in a stream are isotopically distinct, a combination of 13C and 15N analyses can yield considerable insight into a stream food
web. Such an analysis was performed for the dominant
consumers and their food resources of Lookout Creek
in Oregon (Fig. 26; Fry, 1991). In Lookout Creek, the
15
N values revealed that macroinvertebrates were
trophically distinct from two predatory vertebrates
studied, whereas the 13C values showed that consumers
relied more heavily on stream algae than on terrestrial
detritus. The macroinvertebrate scrapers studied (the
snail Juga and the caddisfly Dicosmoecus gilvipes)
appeared to consume algae, whereas the shredding
caddisfly Heteroplectron californicum apparently ate
terrestrial detritus. The stonefly Calineuria californica,
a presumed predator, occupied a trophic level close to
FIGURE 26
Stable isotope cross-plot of selected members of Lookout Creek, Oregon, food web. Animals horizontally positioned over
other sources (i.e., have similar 13C) are inferred to consume those
materials. Animals, displaced vertically 2 – 4 15N units upward are
inferred to belong to the next higher trophic level in the food web.
[Modified from Fry, 1991.]
primary consumers and thus may have consumed some
plant matter along with small invertebrates that were
not measured. The endosymbiotic midge larva Cricotopus nostocicola (described earlier) was positioned over
its algal host Nostoc parmeliodes. Cutthroat trout,
Oncorhynchus clarki, and Pacific giant salamanders,
Dicamptodon ensatus, consumed invertebrates, as
shown by their vertical positioning over the 13C of their
food and a 3 – 4 per mil difference in 15N (Fig. 26). Gut
contents of trout revealed a variety of invertebrates
whereas salamanders had exclusively snails in their
guts (R. Wildman, Oregon State University, personal
communication).
Use of stable isotopes to trace organic matter
sources for insect consumers has been a particularly
powerful tool in streams that support runs of anadromous salmon. Pacific salmon die after spawning, and
the carcasses provide nutrients that are used by stream
algae (e.g., Brickell and Goering, 1970; Mathisen et al.,
1988), which are then fed upon by grazers (e.g., Richey
et al., 1975; Kline et al., 1990). Salmon carcasses also
serve as a high quality source of CPOM for stream
shredders (Kline et al., 1990). Stable isotope technology provides a means to quantify the importance of
salmon carcasses to the stream food web. Salmon returning to streams to spawn have an isotopic signature
that reflects their position in the marine food web.
They are enriched in 15N, reflecting their predation on
marine fishes, and enriched in 13C reflecting their dependence on marine sources of carbon. By comparing
the isotopic signature of algae and insects in reaches
18. Aquatic Insect Ecology
761
TABLE IV 15N and 13C values for food-web components and source materials in a Lake
Superior tributary stream
15
Material
Upstream
Salmon
CPOM
Hydropsyche
Heptageniidae
0.2
3.1
1.6
13
N
Downstream
Upstream
8.9
4.1
4.0
28.5
29.1
29.2
C
Downstream
22.8
28.9
28.9
Salmon spawning occurs in the downstream reach of the river, but not the upstream reach. The stable
isotope value shows that salmon-derived N, but not salmon-derived C, is incorporated into collector
(Hydropsyche) and especially grazer (Heptageniidae) food-web components. [From Schuldt and Hershey, 1995.]
not used by salmon, to reaches where salmon spawn,
the proportion of marine-derived N and C (from
salmon) in the insect community can be traced. Thus,
Kline et al. (1990) showed that at least 90% of the N
in periphyton from Sashin Creek, in southeastern
Alaska, was derived from spawning salmon or salmon
eggs. Herbivores contained 50 – 100% salmon-derived
N. In a Lake Superior tributary stream, the 15N signature of heptageniid grazers suggested about a 30% reliance on salmon-derived N from Lake Superior (Table
IV; Schuldt and Hershey, 1995).
V. SECONDARY PRODUCTION
Secondary production is the amount of animal biomass that is produced in a given area over a given time
period, usually expressed in g / m2 per year. For a cohort, this measurement integrates both individual
growth rate and survivorship of individuals within the
cohort (see Benke, 1984, 1996). In a food-web context,
secondary production is the summation of the amount
of biomass that is available to be consumed by the next
trophic level over some period of time. Thus, an estimate of secondary production expresses energy or material flow through the consumer components of the
ecosystem, and is useful for identifying major pathways
of energy flow in an ecosystem (e.g., Smock and Roeding, 1986; Benke and Wallace, 1997).
Estimation of secondary production often provides
insights that are not apparent from biomass or abundance measurements alone. This is especially true for
organisms of small body size if they exhibit rapid
growth rates. For example, Benke (1998) found extremely high secondary production of snag-dwelling
chironomids in a high-order blackwater river despite
their low biomass. This was due to very high biomass
turnover rates of these small-bodied organisms (Benke,
1998). In Sycamore Creek, Arizona, macroinvertebrate
secondary production, which was primarily insect production, was also extremely high despite fairly low biomass (Fisher and Gray, 1983). This high rate of production reflected a long growing season, small adult
size, very rapid life cycles, and abundant (if low quality) FPOM (Fisher and Gray, 1983). But, even for large
bodied insects measurement of production may provide
surprising insights. O’Hop et al. (1984) showed that
production of the detritivorous stonefly Peltoperla
maria was similar in a disturbed and undisturbed
stream despite more than twofold differences in abundances, reflecting faster growth and higher density of
larger individuals in the disturbed stream. Although
stream secondary production is very often dominated
by insects, crustaceans and molluscs may also be quite
important (see Waters and Hokenstrom, 1980; Fisher
and Gray, 1983). In a recent study of production in a
neotropical stream, Ramírez and Pringle (1998) show
that insect production is low despite rapid growth
rates. They attribute this low production to a combination of high abundance of non-insect macroconsumers
and scouring of resources by flooding.
Because different habitats vary markedly in abundances and community composition of their insect
fauna, calculation of secondary production on a habitat-specific basis is needed to reveal the spatial complexity of insect-mediated pathways of energy flow in
aquatic ecosystems. In addition, failure to consider
some habitats may grossly underestimate secondary
production. For example, Smock et al. (1992) showed
that debris dams were very important compared to
other habitats in one low-gradient first-order stream,
but that the floodplain was most productive in another
stream. In addition, the vertical extent of the hyporheic
zone in the two streams accounted for a large difference in secondary production between the two systems
(Smock et al., 1992). Snag habitats are very important
762
A. E. Hershey and G. A. Lamberti
sites of secondary production in higher order rivers,
reflecting the success of both mayflies and chironomids
in utilizing the amorphous detritus associated with that
habitat (Benke and Jacobi, 1994; Benke, 1998).
Fisheries managers often use secondary production
to assess the capacity of a stream reach to support fish.
In many cases, there does not appear to be enough invertebrate production to support the observed fish production, a problem referred to as Allen’s paradox
(Allen, 1951). This may be due to underestimates of
the production-to-biomass ratio (P / B) for individual
species (Benke, 1996), or perhaps failure to consider
the production of chironomid midges (Huryn and
Wallace, 1986; Berg and Hellenthal, 1992). Recently,
Huryn (1996) has shown that trout can consume a very
large proportion of the benthic macroinvertebrates
(mostly insects) in a stream.
Although conceptually very useful, secondary production is time-consuming and expensive to measure at
the community level. The varied techniques for measuring secondary production, reviewed by Benke (1984,
1996), involve sampling efforts that quantify consumer
standing stocks at several points in the growing season,
at least a trophic level understanding of the various
consumers, and some knowledge of life histories of the
dominant consumer components (Table V). However,
in the process of measurement, considerable insight is
gained into the insect community, which sheds light on
which consumers play key roles in the various aquatic
ecosystem processes.
TABLE V Methods for measuring secondary production of stream macroinvertebrates. Modified from Benke, 1984, 1996,
where secondary production methods are reviewed. [Modified from Hershey and Lamberti, 1998]
Method
Data needed
Comments
Other references
Allen curve
Density (No. / m2) for several
dates spanning development
of a cohort; mean individual
mass for each date [often obtained
from length weight regressions
(Smock, 1980)].
Density (No. / m2) for several dates
spanning development of a cohort;
mean individual mass for each date
[often obtained from length weight
regressions (Smock, 1980)].
Density (No. / m2) for several dates
spanning development of a cohort;
mean individual biomass for each date
[often obtained from length weight
regressions (Smock, 1980)].
Growth rate; mean biomass
Graphical presentation of density versus mean
individual mass. The area under the curve
corresponds to the secondary production of a
population. This method was developed prior
to the widespread availability of electronic
calculators and computers.
Production (P) between sampling dates is calculated
as the product of mean density (N) and change
in mean individual mass (W): P NW.
Annual production is the summation of these
incremental measurements for a year.
Production lost between sampling dates is the
product of the change in density and the
mean individual mass: P WN.
Annual production is the sum of all
production losses over a year.
Production between sampling dates is the
product of growth rate and mean biomass,
where growth rate is determined in laboratory
experiments, or from field samples of distinct
cohorts, and biomass is determined from field
samples: P g B. Growth is temperature and
size-dependent, so accuracy of this method
depends on making measurements under the
appropriate conditions. Annual production
is: P (gi)(Bi), where the I’s are the
production intervals.
This method is widely used and does not depend
on the presence of distinct cohorts in field
populations. The mean size – frequency distribution,
which is skewed toward smaller size categories
due to mortality, represents an average cohort. For
populations where distinct cohorts are known, this
method gives a similar result as the increment
summation method. P WNi, where
i number of size classes. This estimate of
secondary production needs to corrected for
the CPI: (multiply P by 365 / CPI)
Allen, 1951
Increment-summation
Removal-summation
Instantaneous growth
Size frequency
Mean annual density; size distribution
of individuals throughout year; mean
mass for each size category; cohort
production interval (mean development
time from hatching to emergence).
Waters, 1977;
Waters and
Crawford, 1973
Waters, 1977
Ross and Wallace,
1981; Hauer and
Benke, 1987
Benke, 1979
18. Aquatic Insect Ecology
VI. EFFECTS OF LAND USE ON
AQUATIC INSECT COMMUNITIES
Because aquatic insects are sensitive to physical and
chemical conditions in their environment as well as to
top-down and bottom-up biotic controls, their communities also reflect changes in land use on both geological
and ecological time scales and as well as on multiple
spatial scales. For both lotic and lentic ecosystems, nutrient loading from either nonpoint or point sources,
changes in hydrology, timber harvesting, construction,
and riparian or shoreline development are among the
most important land-use alterations that affect insects.
Lentic insect responses to land-use disturbances are
seen through indirect effects of physical and chemical
changes in littoral macrophyte zones and chemical
changes in profundal zones. Nutrient loading results
from many land-use changes, including nutrients associated with sediments derived from construction and
tillage; fertilizer from agriculture, suburban and urban
runoff; and wastewater and industrial inputs (Carpenter et al., 1998). These inputs are derived from both inflow streams and land uses on lake shorelines themselves. Nutrient inputs enhance macrophyte growth
and / or phytoplankton productivity. Since macrophytes
support a high diversity of aquatic insects as well as affording refuge from fish predators (Thorp and Bergey,
1981; Crowder and Cooper, 1982; Gilinsky, 1984;
Hershey, 1985, and see Fig. 22;), land-use changes
altering macrophytes have strong indirect effects on littoral insects. However, if macrophyte growth becomes
too dense, anoxia may be a problem, especially at the
sediment / water interface. Low dissolved oxygen dramatically affects insect community structure (see Section II.B). Sedimentation may negatively affect macrophytes, thereby indirectly negatively affecting insect
abundance and diversity. Nutrient inputs also enhance
phytoplankton production (e. g., Schindler, 1977;
Axler and Reuter, 1996), and at high level this often
leads to bottom-water anoxia and associated low insect
diversity and changes in insect community structure
(see Hilsenhoff, 1966; Saether, 1979, and Section II.B).
Invertebrate populations in wetlands also are affected by vegetational and water management practices
and by water quality and plant physical structure (see
Ferenc et al., 1999). However, Wolf et al. (1999) found
little direct effect of land use on wetland invertebrates
in urban playa wetlands, although they did find invertebrate abundances were positively associated with
macrophyte growth. Similar to littoral areas of lakes,
macrophyte beds respond to land use surrounding the
wetlands. Water-level fluctuation in wetlands, often associated with land-use changes, has considerable influence on invertebrate community structure and the bal-
763
ance between insect and noninsect invertebrates
(Anderson et al., 1999; Hershey et al., 1999). Wetland
insects and other invertebrates are a major source of
food for migratory and resident waterfowl (Swanson,
1984 / 1985; Martin et al., 1951; Tester, 1995), thus
wetland loss and associated loss of invertebrate food
resources and habitat have had a major impact on wetland bird species (Richardson, 1979).
Hynes (1975) first articulated the importance of
employing a landscape perspective for understanding
stream ecosystem structure and function. The realization of this linkage between the stream and its terrestrial setting was a critical development in stream
ecosystem theory (Minshall et al., 1985) and aquatic
insects have been central to this perspective because of
their role in processing allochthonous inputs. This
aquatic – terrestrial relationship is well appreciated
qualitatively, but poorly understood quantitatively.
Local geomorphology, vegetation, and climate determine the quality and quantity of organic matter entering a stream. These processes and disturbance history
at a site determine the spatial and temporal patterns of
riparian zone characteristics that dictate the physical
characteristics of the stream habitat (Gregory et al.,
1991). Thus, riparian processes and natural disturbances, or disturbances due to changes in land use, will
alter insect habitats in the stream channel. These disturbances operate over temporal scales ranging from
geologic alterations of landform, to ecological succession of the riparian forest, as well as over spatial scales
spanning watersheds, to local patchiness of algal
growth on individual rocks within the channel (Gregory et al., 1991). Aquatic insect abundance and diversity responds to all of these perturbations.
Aquatic insect communities reflect land use within
the watershed because changes in land use alter stream
habitat and water quality (Gregory et al., 1987; Chauvet and Décamps, 1989; Merritt and Lawson, 1992).
As discussed previously, insect communities are sensitive to predominant sources of organic matter, dictated
largely by stream order, local geomorphology, and riparian vegetation (Vannote et al., 1980; Naiman et al.,
1987; Naiman and Anderson, 1996). Biogeography
(Resh, 1992; Hershey et al., 1995b) and water quality
factors (Richards and Minshall, 1992; Lamberti and
Berg, 1995) also constrain insect communities.
Use of geographic information systems has facilitated landscape-level analyses and land-use studies of
stream insect communities. In Minnesota north shore
tributaries to Lake Superior, insect community structure
and substrate characteristics within a stream are correlated with agricultural and residential land uses in the
watersheds (Richards and Host, 1994). The types of
alterations seen will depend on the nature and intensity
764
A. E. Hershey and G. A. Lamberti
FIGURE 27
Theoretical relationship between macroinvertebrate species diversity and frequency of disturbance (e.g., floods) in streams ecosystems [redrawn from Ward and Stanford (1983) after Connell (1978)],
with examples of macroinvertebrates that can be found under different conditions in North America. Rarely
disturbed streams often contain large, long-lived taxa such as (top to bottom): Neohermes hellgrammite, Argia
damselfly, and Dicosmoecus caddisfly. Streams with intermediate levels of disturbance usually contain a diverse
fauna including (clockwise from upper left): Calineuria stonefly, Tipula cranefly, Drunella mayfly, Glossosoma
caddisfly, Juga snail, Hydropsyche caddisfly, and Simulium black fly. Streams that are disturbed frequently may
contain small-bodied taxa with short life cycles or high reproductive rates such as (clockwise from upper left):
Helicopsyche caddisflies, Baetis mayflies, and chironomid midges.
of the land-use change (Cairns, 1990), but local geomorphology modifies some of the effects of land-use
change, as well as constraining land cover and dictating
what land uses might occur in a watershed (McIntire
and Colby, 1978; Naiman et al., 1993). In an extensive
study of a 45-catchment river basin in central Michigan,
Richards et al. (1996) found that geology and land use
were approximatlely of equal importance in determining invertebrate communities in streams.
Land-use changes that alter large woody debris
(LWD) have large impacts on insect communities (e.g.,
Fig. 27). Logging is one of the major such land uses.
Logging has greatly altered the size and amount of
LWD that enters stream channels (Sullivan et al.,
1987). Removal of riparian trees changes inputs of
LWD for centuries, altering substrate stability and insect habitat (Naiman and Anderson, 1996); but recent
changes in forest practices now offer some protection
to the riparian zone from excessive harvest (Gregory
and Ashkenas, 1990). Streams that lack LWD sources
due to past logging are candidates for rehabilitation,
which now emphasizes the input of logs in natural volumes and arrangements to improve stream habitat and
to augment the food base. In addition to removing
LWD and thereby altering detrital inputs and channel
structure, logging also removes vegetative cover within
the watershed. This results in increased sedimentation
into the stream, reduced infiltration which decreases
groundwater recharge, and an increased runoff which
increases the magnitude of stream discharge change following storm events (e.g., Gregory et al., 1991;
Naiman and Anderson, 1996).
Other land uses, including urban, agricultural, and
industrial development may follow timber harvest.
18. Aquatic Insect Ecology
Agriculture has a large effect on sedimentation in a
stream reach (Allan et al., 1997), but stream buffers are
more important than the whole catchment in determining sediment-related habitat variables in agricultural
basins (Richards et al., 1996). Road construction is
usually associated with all types of human land use.
Road construction often destabilizes hillsides, leading
to landslides and sometimes debris flows in stream
channels downslope (Lamberti et al., 1991; Swanson
et al., 1987), which constitute a major disturbance to
the stream ecosystem (see below). These changes in
land use dramatically alter watershed hydrology, and
hence stream hydrology. As discussed previously, insect
communities are strongly affected by stream hydrology,
but are also sensitive to the various forms of pollution
that result from many land uses (see Richards et al.,
1993). Thus, insect communities are intimately linked
to land use in the watershed as well as to local conditions in the stream channel.
VII. ROLE OF DISTURBANCE
A “disturbance” has been defined as any discrete
event that disrupts population, community, or ecosystem structure, usually by changing resource abundance
or the physical environment (Sousa, 1984; Resh et al.,
1988). Many types of natural and human-induced disturbance can disrupt the habitat, resources, or population densities of aquatic insects. Large-scale natural disturbances that can affect insects include: (1) hydrologic
events such as floods, ice, or wave action that scour or
remove substrates; (2) desiccation events such as
droughts that lead to drying of aquatic habitats; (3)
watershed disturbances, such as wildfire, that can disrupt nutrient and sediment inputs to streams and lakes
for decades; and (4) seasonal events, such as summer or
winter oxygen depletion, that can lead to insect mortality and major changes in community structure (see Fig.
27). Naturally, many small-scale disturbances can also
affect local insect populations. Human-related disturbances of aquatic insect habitats are at least as important and include nominally: (1) point and nonpoint
source pollutants that enter aquatic ecosystems from
the land, connected water, or the atmosphere; (2) water
withdrawal, diversion, and storage; (3) modifications
of river channel geomorphology and destruction of
wetlands; (4) watershed disruptions, such as logging
and other land uses; and (5) introduction and establishment of exotic species. All of these mechanisms, singly
and in combination, can alter population densities and
community structure of insects, sometimes for decades
or centuries as in the case of watershed land-use
change. In general, however, natural disturbances tend
765
to be episodic and, if the habitat returns to its orginal
state, insect communities recover quickly. Human disturbance, by contrast, tends to be chronic and thus insect communities may not recover until the disturbance
ceases or the habitat is restored (Cairns, 1990).
Variation in flow (floods to desiccation) is the
major cause of natural disturbance in streams and is responsible for large, usually temporary, reductions in
insect abundance and diversity (Fisher et al., 1982;
Lamberti et al., 1991). Stream insects have evolved
various mechanisms to deal with such flow variation,
including life-history adjustments to minimize the presence of vulnerable stages during times of peak flows,
behavioral movement into more protected hyporheic
(interstitial) and lateral habitats during floods, and
high reproductive rates to compensate for losses. Despite these adaptations, severe flood disturbance will
still remove large proportions of the insect fauna. During drought, refuge provided by the deep hyporheic
zone is especially important, but its accessibility to insects will depend on its grain size. Other insects may
survive drought via resistant life stages, such as diapausing eggs or cocoons. Insects can recolonize streams
from several sources: (1) egg laying by adults, usually
aerial in the case of insects; (2) drift from upstream
areas; and (3) movement along the bed from upstream,
downstream, hyporheic, or lateral habitats (Smock,
1996). These behaviors are discussed in more detail
above (see Section II. D. 3)
In general, aquatic insect communities recover
rapidly from natural disturbance. Lamberti et al.
(1991) studied the response and recovery of macroinvertebrates (mostly insects) in a Cascade Mountain
stream that experienced an extreme disturbance. A debris flow, which is an episodic mass movement of sediment and debris through the stream channel (usually
triggered by a landslide), devastated about a 1-km
reach of Quartz Creek, Oregon, and partially disturbed
areas downstream (Fig. 28). More than 99% of the
macroinvertebrates were removed by the debris flow.
An associated flood alone in an upstream reach removed 90% of the invertebrates, so that in-stream
sources of insects clearly were low. Yet, in both the
flood-impacted and debris flow-impacted reaches,
macroinvertebrate species richness and total abundance
recovered within one year of the disturbance to “normal” levels (Fig. 28). In another example, the eruption
of Mt. St. Helens (southwest Washington) eliminated
the entire biota from entire drainage systems, which included streams, lakes, and wetlands. Many stream
channels were completely rerouted by topographic
changes. Yet, in the first year after the eruption, 98
species of macroinvertebrates (mostly insects) were
found in one devastated stream (Anderson, 1992). Five
766
A. E. Hershey and G. A. Lamberti
FIGURE 28
Debris flow and its effects in Quartz Creek, Oregon. (A) Upstream reach that was flooded only.
(B) Reach most severely impacted by debris flow of February 1986. (C) Downstream of debris flow, showing
debris dam and partial impact. (D) Recovery of periphyton chlorophyll a in the three reaches. (E) Recovery of
macroinvertebrate density (mostly insects). (F) Recovery of macroinvertebrate species richness. [Modified from
Lamberti et al., 1991.]
years after the eruption, 141 species were found in the
same stream, although nearly half of these were the
ubiquitous chironomid midges. Both of these examples
attest to the remarkable resilience of stream insects and
other invertebrates to natural disturbance.
Episodic natural events contrast, however, with
chronic anthropogenic disturbance, such as inputs of
chemical and thermal pollutants or prolonged watershed
disturbance. Prolonged point-source pollution can have
devastating effects on aquatic insects. Wallace and Brady
(1971) reported that low concentrations (0.2 ppm) of
the insecticide dieldrin in a factory effluent reduced benthic insect densities by over 90% in a South Carolina
stream, and that the remaining invertebrates had high
tissue concentrations (up to 100 ppm) of dieldrin. Wallace and coworkers later explored the structural and
18. Aquatic Insect Ecology
functional significance of reduced insect populations in
streams by conducting an experimental release of insecticide (methoxychlor) in an Appalachian Mountain
stream. Insecticide application resulted in massive drift
of insects, and reduction of benthic densities by over
85% (Wallace et al., 1982, 1989). The remaining benthic invertebrates were composed of an impoverished
fauna of mostly noninsects (e.g., oligochaetes) and chironomid midge larvae. Furthermore, important ecological processes in the stream were altered. For example,
leaf litter decomposition declined drastically because an
important shredding caddisfly, Lepidostoma, was eliminated by the insecticide treatment (Whiles et al., 1993).
When treatment ceased, Lepidostoma recolonized
rapidly and leaf litter processing recommenced, again
pointing to the resilience of aquatic insect communities.
However, human chronic disturbance of aquatic ecosystems and their watersheds tends to preclude recovery of
insect communities to predisturbance conditions. Watershed disturbances such as forest logging, agriculture, and
urbanization can have long-term impacts on aquatic insect communities by altering energy inputs, thermal
regimes, and sediment composition (Gregory et al.,
1987; Lamberti and Berg, 1995; Wallace et al., 1988).
VIII. AQUATIC INSECTS IN
BIOMONITORING STUDIES
Biomonitoring studies are used to measure response and recovery of aquatic communities to disturbances, protect biodiversity, evaluate compliance, and
improve understanding of the relationship between
physical, chemical, and biological components (Gurtz,
1994). Many biomonitoring programs incorporate fish,
macroinvertebrate, and algae components in their studies, although macroinvertebrate studies, especially
those focusing on aquatic insects, are widely used by
many agencies. The National Water Quality Assessment (NAWQA) program of the USGS seeks to evaluate water quality at spatial scales ranging from local to
national (Gurtz, 1994). Such biomonitoring studies are
used to assess aquatic ecosystem health because organisms function as sensors of the quality of their environment in several ways that direct measurements of water
quality cannot (Loeb, 1994). Insects have been particularly useful in biomonitoring of streams but also are
used in lentic studies (Brinkhurst, 1974; Warwick and
Tisdale, 1989) to assess changes in lake water quality
and productivity. Warwick (1980) also has used insects
in paleolimnological studies to reconstruct changes in
land use surrounding a lake.
Stream macroinvertebrates, especially insects, offer
distinct advantages in biomonitoring studies (Stewart
767
and Loar, 1994). Their small size and limited mobility
renders them relatively easy to sample. They also are
known to be sensitive to a variety of pollutants. For example, insects absorb or transform chemical pollutants,
such that direct chemical analyses may not be measuring toxicity in the same way as do insects. Insects also
integrate environmental quality over a longer time interval than do static measures of water quality. Thus, biomonitoring incorporates and integrates ecology, life history, physiology, and taxonomy into a management and
assessment tool. Although valuable and widely used,
there are also a few disadvantages to using aquatic insects in these studies (Stewart and Loar, 1994). Spatial
and temporal variation in abundance of aquatic life-history stages must be accounted for. Thus, an appropriate
sampling design in a biomonitoring program must stratify this spatial and temporal variability to account for
as much of it as possible (Hughes et al., 1994). Stratification according to underlying geology is also of critical
importance (Minshall et al., 1985; Richards et al.,
1996). Furthermore, aquatic insect identification is
time-consuming and requires trained personnel; errors
in taxonomy can lead to erroneous conclusions (see
Lenat and Barbour, 1994; Rosenberg and Resh, 1996).
Rosenberg and Resh (1996) provided a comprehensive review of current biomonitoring approaches
that use aquatic insects. These approaches span a range
of scales: (1) biochemical and physiological measurements, including metabolic studies and measures of enzyme activities; (2) individual attributes such as morphological, behavioral, and life-history parameters, or
use of “sentinel organisms” that serve as bioaccumulators of toxic materials; (3) population- and assemblagelevel responses, such as using occurrence or abundance
of indicator species as a measure of sensitivity to a pollutant; (4) community-level approaches which synthesize many types of data into summary responses to a
pollutant; and (5) ecosystem-level scales that assess effects of stressors on processes and function.
Of these, community-level approaches are probably
in most widespread use, and these may be either qualitative or quantitative. Rapid assessment methods are often based on insect communities, are generally qualitative, but permit assessment of many streams with a
reasonable degree of effort (see Lenat and Barbour,
1994). However, quantitative approaches are also very
common, and the choice between qualitative and quantitative techniques depends on the objectives of the
study. Several different metrics of aquatic insect communities are used (Rosenberg and Resh, 1996). Taxa
richness is widely used, but is difficult to compare
across studies because different levels of resolution tend
to be used by different investigators and for different
taxonomic groups. EPT richness, which is the combined
768
A. E. Hershey and G. A. Lamberti
an integrated picture of the recent history of an aquatic
ecosystem.
IX. SUMMARY
FIGURE 29
Richness of EPT (Ephemeroptera, Plecoptera, and Trichoptera) taxa is robust to seasonal variation within a site, although
it varies longitudinally and between streams. [From Stewart and
Loar, 1994.]
richness of Ephemeroptera, Plecoptera, and Trichoptera
(EPT taxa) takes advantage of the fact that EPT taxa
are relatively easy to identify and are pollution-sensitive
(Lenat, 1988; Plafkin et al., 1989; Resh et al., 1996).
This index varies longitudinally but is robust to seasonal and interannual fluctuations (Fig. 29, Stewart and
Loar, 1994). Because chironomids as a group are considered pollution-tolerant and EPT taxa are not, the ratio of EPT / chironomids is another popular communitylevel metric. Diversity and similarity indexes afford the
advantage of being readily comparable between sites,
and biotic indexes offer a readily interpretable score
(e.g., Hilsenhoff, 1982). Functional feeding group measures rely on functional rather than taxonomic information, and in this sense they bridge community- and
ecosystem-level approaches to biomonitoring.
Rather than relying on a single metric for assessment in biomonitoring studies, multimetric and multivariate approaches often are recommended (ter Braak
and Prentice, 1988; Karr, 1994; Lenat and Barbour,
1994; Richards et al., 1996; Rosenberg and Resh,
1996). Multimetric approaches can combine the
strengths of individual metrics to minimize the chance
of an incorrect assessment, and often several metrics
can be calculated from the same data set. Multivariate
approaches permit partitioning of variance due to different factors, thus providing more useful information
than a single measure. They also are less subjective
than are univariate approaches. Resh et al. (1996) provide a useful exercise that demonstrates the utility of
several different biomonitoring approaches that use
benthic macroinvertebrates. They stress the importance
of combining information on both physical habitat
quality and macroinvertebrate composition when assessing water quality. A cornerstone of their approach
is that macroinvertebrates, especially insects, provide
The study of aquatic insect ecology is dynamic,
with several hundred new papers being added to the literature each year. While it is not possible to provide a
comprehensive treatment of aquatic insect ecology in a
single chapter, we provide a broad overview of major
topics in aquatic insect ecology. Our specific goals were
to: (1) describe the basic life histories of aquatic insects
and their general community structure; (2) outline their
role in major ecological processes involving insects that
occur in aquatic ecosystems; (3) discuss how the structure of insect communities and the services they perform are affected by disturbance to their aquatic
ecosystems and the changing landscape in which they
are found; and (4) summarize how their communities
can be used by managers to assess aquatic ecosystem
health. This synopsis should provide a useful starting
point for those seeking an entry into the rich literature
on aquatic insect ecology.
LITERATURE CITED
Allan, J. D. 1982. The effects of reduction in trout density on the invertebrate community of a mountain stream. Ecology
63:1444 – 1455.
Allan, J. D. 1995. Stream ecology, Chapman & Hall, London., 388 p.
Allan, J. D., Erickson, D. L., Fay, J. 1997. The influence of catchment
land us on stram integrity across multiple spatial scales. Freshwater Biology 37:149 – 161.
Allan, J. D., Feifarek, B. P. 1989. Distances travelled by drifting
mayfly nymphs: factors influencing return to the substrate. Journal of the North American Benthological Society 8:322 – 330.
Allen, K. R. 1951. The Horokiwi Stream. A study of a trout population. New Zealand Department of Fisheries Bulletin 10:1 – 238.
Anderson, N. H. 1976a. Carnivory by an aquatic detritivore, Clistoronia magnifica (Trichoptera: Limnephilidae). Ecology
57:1981 – 1085.
Anderson, N. H. 1976b. The distribution and biology of the Oregon
Trichoptera. Oregon State University Agricultural Experiment
Station Technical Bulletin 134, Corvallis, OR, 152 p.
Anderson, N. H. 1992. Influence of disturbance on insect communities in Pacific Northwest streams. Hydrobiologia 248:79 – 92.
Anderson, N. H., Bourne, J. R. 1974. Bionomics of three species of
glossosomatid caddisflies (Trichoptera: Glossosomatidae) in
Oregon. Canadian Journal of Zoology 52:405 – 411.
Anderson, N. H., Hanson, B. P. 1987. An annotated check list of
aquatic insects collected at Berry Creek, Benton County, Oregon
1960 – 1984. Occasional Publication Number 2, Systematic Entomology Laboratory, Department of Entomology, Oregon
State University.
Anderson, N. H., Sedell, J. R. 1979. Detritus processing by macroinvertebrates in stream ecosystems. Annual Review of Entomology 24:351 – 377.
18. Aquatic Insect Ecology
Axler, R. P., Reuter, J. E. 1996. Nitrate uptake by phytoplankton and
periphyton: Whole-lake enrichments and mesocosm 15N experiments in an oligotrophic lake. Limnology and Oceanography
41:659 – 671.
Barnes, H. T. 1928. Ice engineering, Renouf Pub. Co., Montreal.,
364 p.
Batzer, D. P., Resh, V. H. 1996. Trophic interactions among a beetle
predator, a chironomid grazer, and periphyton in a seasonal
wetland. Oikos 60:251 – 257.
Batzer, D. P., Wissinger, S. A. 1996. Ecology of insect communities in
nontidal wetlands. Annual Review of Entomology 41:75 – 100.
Beeton, A. M. 1961. Environmental changes in Lake Erie. Transactions of the American Fisheries Society 87:73 – 79.
Benke, A. C. 1979. A modification of the Hynes method for estimating
secondary production with particular significance for multivoltine
populations. Limnology and Oceanography 24:168 – 174.
Benke, A. C. 1984. Secondary production of aquatic insects, in Resh,
V. H., Rosenberg, D. M., Eds., The ecology of aquatic insects.
Praeger, New York, pp. 289 – 322.
Benke, A. C. 1993. Concepts and patterns of invertebrate produciton
in running waters. Internationale Verinigung fur theoretische
und angewandte Limnologie, Verhandlungen 25:15 – 38.
Benke, A. C. 1996. Secondary production of macroinvertebrates, in
Hauer, F. R., Lamberti, G. A., Eds., Methods in stream ecology.
Academic Press, San Diego, CA, pp. 557 – 578.
Benke, A. C. 1998. Production dynamics of riverine chironomids: extremely high biomass turnover rates of primary consumers.
Ecology 79:899 – 910.
Benke, A. C., Wallace, J. B. 1980. Trophic basis of production
among net-spinning caddisflies in a southern Appalachian
stream. Ecology 61:108 – 118.
Benke, A. C., Van Arsdall, T. C., Jr., Gillespie, D. M., Parrish, F. K.
1984. Invertebrate poductivity in a subtropical blackwater
river: the importance of habitat and life hitory. Ecological
Monographs 54:25 – 63.
Benke, A. C., Hunter, R. J., Parrish, F. K. 1986. Invertebrate drift dynamics in a subtropical blackwater river. Journal of the North
American Benthological Society 5:173 – 190.
Benke, A. C., Jacobi, D. I. 1994. Production dynamics and resource
utilization of snag-dwelling mayflies in a blackwater river. Ecology 75:1219 – 1232.
Benke, A. C., Wallace, J. B. 1997. Trophic basis of production
among riverine caddisflies: implications for food web analyses.
Ecology 78:1132 – 1145.
Berg, M. B., Hellenthal, R. A. 1992. Role of Chironomidae in energy
flow of a lotic ecosystem. Netherlands Journal of Aquatic Ecology 26:471 – 476.
Bergey, E. A., Resh, V. H. 1994. Interactions between a stream caddisfly and the algae on its case: factors affecting algal quantity.
Freshwater Biology 31:153 – 163.
Bilby, R. E., Ward, J. W. 1989. Changes in characteristics and function
of woody debris with increasing stream size in western Washington. Transactions of American Fisheries Society 118:368 – 378.
Bird, G. A., Hynes, H. B. N. 1981. Movement of immature insects in
a lotic habitat. Hydrobiologia 77:103 – 112.
Bisson, P. A., Bilby, R. E., Bryant, M. D., Dolloff, C. A., Brette, G. B.,
House, R. A., Murphy, M. L., Koski, K. V., Sedell, J. R. 1987.
Large woody debris in forested streams in the Pacific Northwest: past, present, and future, in Salo, E. O., Cundy, T. W.,
Eds., Streamside management: forestry and fishery interactions.
University of Washington Institute of Forest Resources Contribution No. 57, pp. 143 – 190.
Bohannan, R. E., Johnson, D. M. 1983. Response of littoral invertebrate populations to a spring fish exclusion experiment. Freshwater Invertebrate Biology 2:28 – 40.
769
Borror, D. J., Triplehorn, C. A., Johnson, N. F. 1989. An
introduction to the study of insects, 6th ed., Saunders, Philadelphia, PA.
Brickell, D. C., Goering, J. J. 1970. Chemical effects of salmon decomposition on aquatic ecosystems, in Murphey, R. S., Ed. First
International Symposium of Water Pollution Control in Cold
Climates. U. S. Government Printing Office, Washington, DC,
pp. 125 – 138.
Brinkhurst, R. O. 1974. The benthos of lakes. St. Martin’s press,
New York, NY.
Brock, E. M. 1960. Mutualism between the midge Cricotopus and
the alga Nostoc. Ecology 41:474 – 483.
Brock, T. D. 1978. Thermophilic microorganisms and life at high
temperatures, Springer, New York, NY. 465 p.
Brown, K. S. 1996. Do disease cycles follow changes in the weather?
BioScience 46:479 – 481.
Brundin, L. 1949. Chironomide und andere Bodentiere der sudschwedischen Urgebirgsseen. Rep. Inst. Freshwat. Res. Drottningholm 30:1 – 94.
Butler, M. G. 1982. A 7-year life cycle for two Chironomus species in
arctic Alaskan tundra ponds (Diptera: Chironomidae). Canadian Jounral of Zoology 60:58 – 70.
Butler, M. G. 1984. Life histories of aquatic insects, in Resh, V. H.,
Rosenberg, D. M., Eds., The ecology of aquatic insects. Praeger,
New York, NY, pp. 24 – 55.
Cairns, J. Jr. 1990. Lack of theoretical basis for predicting rate
and pathways of recovery. Environmental Management 14:
517 – 526.
Cantrall, I. J., Brusven, M. A. 1996. Semiaquatic Orthoptera, in
Merritt, R. W., Cummins, K. W., Eds., 1996. An introduction to
the aquatic insects of North America, 3rd ed. Kendall / Hunt,
Dubuque, IA, pp. 212 – 216.
Carpenter, S. R. 1982. Stemflow chemistry: effects on population dynamics of detritivorous mosquitoes in tree-hole ecosystems. Oecologia 53:1 – 6.
Carpenter, S. R. 1983. Resource limitation of larval treehole mosquitoes subsisting on beech detritus. Ecology 64:219 – 223.
Carpenter, S. R., Kitchell, J. F., Hodgson, J. R. 1985. Cascading
trophic interactions and lake productivity. BioScience 35:
634 – 639.
Carpenter, S. R., Caraco, N., Correll, D., Howarth, R., Sharpley, A.,
Smith, V. 1998. Nonpoint pollution of surface waters with
phosphorus and nitrogen. Issues in Ecology No. 1. Ecological
Society of America.
Chapman, D. W., Demory, R. L. 1963. Seasonal changes in the food
ingested by aquatic insect larvae and nymphs in two Oregon
streams. Ecology 44:140 – 146.
Chauvet, E., Décamps, H. 1989. Lateral interaction in a fluvial landscape: the River Garonne, France. Journal of the North American Benthological Society 8:9 – 17.
Closs, G. P., Lake, P. S. 1994. Spatial and temporal variation in the
structure of an intermediate stream food web. Ecological
Monographs 64:1 – 21.
Connell, J. H. 1978. Diversity in tropical rain forests and coral reefs.
Science 199:1302 – 1310.
Cooper, S. D., Walde, S. J., Peckarsky, B. L. 1990. Prey exchange
rates and the impact of predation in streams. Ecology 71:
1503 – 1514.
Couch, C. A., Meyer, J. L., Hall, R. O. 1996. Incorporation of bacterial extracellular polymer by blackfly larvae (Simuliidae). Journal of the North American Benthological Society 15:289 – 299.
Crawley, M. J. 1983. Herbivory. The dynamics of animal – plant interactions, Univ. of California Press, Berkeley, CA., 437 p.
Cronin, G., Wissing, K. D., Lodge, D. M. 1998. Comparative feeding
selectivity of herbivorous insects on water lilies: aquatic vs
770
A. E. Hershey and G. A. Lamberti
semi-terrestrial insects and submersed vs floating leaves. Freshwater Biology 39:243 – 57.
Crowder, L. B., Cooper W. E. 1982. Habitat structural complexity
and the interaction between bluegills and their prey. Ecology
63:1802 – 1813.
Cummins, K. W. 1973. Trophic relations of aquatic insects. Annual
Review of Entomology 18:183 – 206.
Cummins, K. W. 1974. Structure and function of stream ecosystems.
BioScience 24:631 – 641.
Cummins, K. W., Klug, M. J. 1979. Feeding ecology of stream invertebrates. Annual Review of Ecology and Systematics 10:147 – 172.
Cummins, K. W., Merritt, R. W. 1996. Ecology and distribution of
aquatic insects, in Merritt, R. W., Cummins, K. W., Eds., An introduction to the aquatic insects of North America, 3rd ed.
Kendall / Hunt, Dubuque, IA, pp. 74 – 86.
Cummins, K. W., Wilzbach, M. A., Gates, D. M., Perry, J. B., Taliaferro, W. B. 1989. Shredders and riparian vegetation. BioScience
39:24 – 30.
Dahm, C. N. 1981. Pathways and mechanisms for removal of dissolved organic carbon from leaf leachate in streams. Canadian
Journal of Fisheries and Aquatic Sciences 38:68 – 76.
Danks, H. V. 1978. Modes of seasonal adaptations in the insects. I.
Winter survival. Canadian Entomologist 110:1167 – 1205.
Deegan, L. A., Peterson, B. J. 1992. Whole river fertilization stimulates fish production in an arctic tundra river. Canadian Journal
of Fisheries and Aquatic Sciences 54:269 – 283.
Delucchi, C. M. 1989. Movement patterns of invertebrates in temporary and permanent streams. Oecologia 78:199 – 207.
Dodds, W. K. 1989. Photosynthesis of two morphologies of Nostoc
parmelioides (Cyanobacteria) as related to current velocities
and diffusion patterns. Journal of Phycology 25:258 – 262.
Dodds, W. K., Marra, J. L. 1989. Behaviors of the midge, Cricotopus
(Diptera: Chironomidae), related to mutualism with Nostoc
parmelioides (Cyanobacteria). Aquatic Insects 11:201 – 208.
Dudley, T. L., Anderson, N. H. 1982. A survey of invertebrates associated with wood debris in aquatic habitats. Melanderia
39:1 – 21.
Duffy, W. G., Liston, C. R. 1985. Survival following exposure to subzero temperatures and respiration in cold acclimatized larvae of
Enallagma boreale (Odonata: Zygoptera). Freshwater Invertebrate Biology 4:1 – 7.
Eriksen, C. H., Resh, V. H., Lamberti, G. A. 1996. Aquatic insect
respiration. in Merritt, R. W., Cummins, K. W., Eds. An introduction to the aquatic insects of North America, 3rd ed.
Kendall / Hunt, Dubuque, IA, pp. 29 – 40.
Feminella, J. W., Hawkins, C. P. 1995. Interactions between stream
herbivores and periphyton: a quantitative analysis of past experiments. Journal of the North American Benthological Society
14:465 – 509.
Fisher, S. G., Gray, L. J. 1983. Secondary production and organic
matter processing by collector macroinvertebrates in a desert
stream. Ecology 64:1217 – 1224.
Fisher, S. G., Gray, L. J., Grimm, N. B., Busch, D. E. 1982. Temporal
succession in a desert stream ecosystem following flash flooding.
Ecological Monographs 52:93 – 110.
Fisher, S. G., Likens, G. E. 1973. Energy flow in Bear Brook, New
Hampshire: an integrative approach to stream ecosystem metabolism. Ecological Monographs 43:421 – 439.
Fry, B. 1991. Stable isotope diagrams of freshwater food webs. Ecology 72:2293 – 2297.
Fry, B., Sherr, E. 1984. 13C measurements as indicators of carbon
flow in marine and freshwater ecosystems. Contributions in
Marine Science 27:13 – 47.
Gallepp, G. W. 1979. Chironomid influence on phosphorus release in
sediment – water microcosms. Ecology 60:547 – 556.
Gilinsky, E. 1984. The role of predation and spatial heterogeneity
in determining benthic community structure. Ecology 65:
455 – 468.
Glime, J. M., Clemons, R. M. 1972. Species diversity of stream insects on Fontinalis spp. compared to diversity on artificial substrates. Ecology 53:458 – 464.
Gotceitas, V., Clifford, H. F. 1983. The life history of Dicosmoecus
atripes (Hagen) (Limnephilidae: Trichoptera) in a Rocky Mountain stream of Alberta, Canada. Canadian Journal of Zoology
61:586 – 596.
Gregory, S. V. 1983. Plant – herbivore interactions in stream systems.
in Barnes, J. R., Minshall, G. W., Eds. Stream ecology. Plenum,
New York, NY, pp. 157 – 189.
Gregory, S. V., Ashkenas, L. R. 1990. Field guide for riparian management. Willamette National Forest. U.S.D.A. Forest Service,
Pacific Northwest Region, 65 pp.
Gregory, S. V., Lamberti, G. A., Erman, D. C., Koski, K. V., Murphy,
M. L., Sedell, J. R. 1987. Influence of forest practices on aquatic
production, in Salo, E. O., Cundy, T. W., Eds., Streamside management: forestry and fishery interactions. University of Washington Institute of Forest Resources Contribution No. 57, pp.
233 – 255.
Gregory, S. V., Swanson, F. J., McKee, W. A., Cummins, K. W. 1991.
An ecosystem perspective of riparian zones. BioScience
41:540 – 552.
Grubaugh, J. W., Wallace, J. B., Houston, E. S. 1997. Production of
benthic macroinvertebrate communities along a southern Appalachian river continuum. Freshwater Biology 37:581 – 96.
Gurtz, M. E. 1994. Design considerations for bioogical components
of the National Water-Quality Assessment (NAWQA) Program,
in Loeb, S. L., Spacie, A., Eds., Biological monitoring of aquatic
systems. Lewis Pub., Boca Raton, FL, pp. 323 – 356.
Hall, R. J., Driscoll, C. T., Likens, G. E. 1987. Importance of hydrogen ions and aluminum in regulating the structure and function
of stream ecosystems: an experimental test. Freshwater Biology
18:17 – 43.
Hart, D. D., Resh, V. H. 1980. Movement patterns and foraging
ecology of a stream caddisfly larva. Canadian Journal of Zoology 58:1174 – 1185.
Hauer, F. R., Benke, A. C. 1987. Influence of temperature and river
hydrograph on blackfly growth rates in a subtropical blackwater river. Journal of the North American Benthological Society
6:251 – 261.
Hauer, F. R., Hill, W. R. 1996. Temperature, light, and oxygen, in
Hauer, F. R., Lamberti, G. A., Eds., Methods in stream ecology.
Academic Press, San Diego, CA, pp. 93 – 106.
Hawkins, C. P., Furnish, J. K. 1987. Are snails important competitors in stream ecosystems? Oikos 49:209 – 220.
Hawkins, C. P., Murphy, M. L., Anderson, N. H. 1982. Effects of
canopy, substrate composition, and gradient on the structure of
macroinvertebrate communities in Cascade Range streams of
Oregon. Ecology 63:1840 – 1856.
Hawkins, C. P., Sedell, J. R. 1981. Longitudinal and seasonal
changes in functional organization of macroinvertebrate communities in four Oregon streams. Ecology 62:387 – 397.
Hayden, W., Clifford, H. F. 1974. Seasonal movements of the mayfly
Leptophlebia cupida (Say) in a brown-water stream of Alberta,
Canada. American Midland Naturalist 91:90 – 102.
Hershey, A. E. 1985. Effects of predatory sculpin on the chironomid
communities in an arctic lake. Ecology 66:1131 – 1138.
Hershey, A. E. 1986. Selective predation by Procladius in an arctic
Alaskan Lake. Canadian Journal of Fisheries and Aquatic Sciences. 43(12) 2523 – 2528.
Hershey, A. E. 1987. Tubes and foraging behavior in larval Chironomidae: implications for predator avoidance. Oecologia 73:236 – 241.
18. Aquatic Insect Ecology
Hershey, A. E., Dodson, S. I. 1987. Predator avoidance by Cricotopus: cyclomorphosis and the importance of being big and hairy.
Ecology 68:913 – 920.
Hershey, A. E, Hiltner, A. L., Hullar, M. A. J., Miller, M. C., Vestal,
J. R., Lock, M. A., Rundle, S., Peterson, B. J. 1988. Nutrient influence on a stream grazer: Orthocladius microcommunities
track nutrient input. Ecology 69:1383 – 1392.
Hershey, A. E., Pastor, J., Peterson, B. J., Kling, G. W. 1993. Stable
isotopes resolve the drift paradox for Baetis mayflies in an arctic river. Ecology 74:2315 – 2325.
Hershey, A. E., Shannon, L., Axler, R., Ernst, C., Mickelson. P.,
1995a. Effects of methoprene and Bti (Bacillus thuringiensis var
israelensis) on non-target insects. Hydrobiologia 308:219 – 227.
Hershey, A. E., Merritt, R. W., Miller, M. C. 1995b. Trophic dynamics, diversity, and life history features of blackflies (Diptera:
Simuliidae) in arctic Alaskan streams, in Chapin, S. F., Kroner,
C., Eds., Arctic and alpine biodiversity. Springer, New York, pp.
283 – 295.
Hershey, A. E., Merritt, R. W., Miller, M. C., McCrea, J. S. 1996.
Organic matter processing by larval black flies in a temperate
woodland stream. Oikos 75:524 – 532.
Hershey, A. E., Peterson, B. J. 1996. Stream food webs, in Hauer,
F. R., Lamberti, G. A., Eds., Methods in Stream Ecology. Academic Press, San Diego, CA, pp. 511 – 530.
Hershey, A. E., Bowden, W. B., Deegan, L. A., Hobbie, J. E., Peterson, B. J., Kipphut, G. W., Kling, G. W., Lock, M. A., Merritt,
R. W., Miller, M. C., Vestal, J. R., Schuldt, J. A. 1997. The Kuparuk River: a long-term study of biological and chemical
processes in an Arctic river, in Milner, A. M., Oswood, M. W.,
Eds., Freshwaters of Alaska. Springer, New York, pp. 107 – 129.
Hershey, A. E., Lamberti, G. A. 1998. Stream macroinvertebrate
communities, in Bilby, R. E., Naiman, R. J., Eds., Ecology and
management of streams and rivers in the Pacific Northwest
coastal regions. Springer, New York, pp. 169 – 199.
Hershey, A. E., Lima, A. R., Niemi, G. J., Regal, R. R. 1998. Effects
of methoprene and Bti (Bacillus thuringiensis var. israelensis) on
non-target macroinvertebrates in Minnesota wetlands. Ecological Applications 8:41 – 60.
Hershey, A. E., Shannon, L., Regal, R., Lima, A., Niemi, G. 1999.
Prairie wetlands of southcentral Minnesota: Effects of drought
on invertebrate communities, in Batzer, D., Wissinger, S., Eds.,
Invertebrates in freshwater wetlands of North America: Ecology
and management. John Wiley and Sons, pp. 515 – 541.
Hilsenhoff, W. L. 1966. The biology of Chironomus plumosus
(Diptera: Chironomidae) in Lake Winnebago, Wisconsin. Annals of the Entomological Society of America 59:465 – 473.
Hilsenhoff, W. L. 1982. Using a biotic index to evaluate water quality in streams. Wisconsin Department of Natural Resources
Technical Bulletin 132:1 – 22.
Hiltner, A. L., Hershey, A. E. 1992. Black fly (Diptera: Simuliidae) response to phosphorus enrichment in an arctic tundra stream.
Hydrobiologia 240:259 – 265.
Hinterleitner-Anderson, D., Hershey, A. E., Schuldt, J. A. 1992. The
effects of river fertilization on mayfly (Baetis sp.) drift patterns
and population density in an arctic river. Hydrobiologia
240:247 – 258.
Hughes, R. M., Heiskary, S. A., Matthews, W. J., Yoder, C. O. 1994.
Use of ecoregions in biological monitoring, in Loeb, S. L., Spacie, A., Eds., Biological monitoring of aquatic systems. Lewis
Pub., Boca Raton, FL, pp. 125 – 154.
Huryn, A. D. 1996. An appraisal of the Allen paradox in a New
Zealand trout stream. Limnology and Oceanography 41:
243 – 352.
Huryn, A. D., Wallace, J. B. 1986. A method for obtaining in situ
growth rates of larval Chironomidae (Diptera) and its applica-
771
tion to studies of secondary production. Limnology and
Oceanography 31:216 – 222.
Hynes, H. B. N. 1970. The ecology of running waters. Univ. of
Toronto Press, Toronto, Ontario. 555 p.
Hynes, H. B. N. 1975. The stream and its valley. Internationale
verein für Theoretische und Angewandte Limnologie Verhandlungen 19:1 – 15.
Jacobs-Lorena, M., Lemos, F. J. A. 1995. Immunological strategies
for control of insect disease vectors: a critical assessment. Parasitology Today 11:144 – 147.
Johnson, D. M., Martin, T. H., Crowley, P. H., Crowder, L. B. 1996.
Link strength in lake littoral food webs: net effects of small sunfish and larval dragonflies. Journal of the North American Benthological Society 15:271 – 288.
Junger, M., Planas, D. 1994. Quantitative use of stable carbon isotope analysis to determine the trophic basis of invertebrate communities in a boreal forest lotic system. Canadian Journal of
Fisheries and Aquatic Sciences 51:52 – 61.
Kajak, Z. 1980. Role of invertebrate predators (mainly Procladius) in
benthos, in Murray, D., Ed. Chironomidae: ecology, systematics,
cytology, and physiology. Pergamon, New York, pp. 339 – 348.
Karr, J. R. 1994. Biological monitoring: Challenges for the future, in
Loeb, S. L., Spacie, A., Eds., Biological monitoring of aquatic
systems. Lewis Pubs., Boca Raton, FL, pp. 357 – 373.
Kidwell, M. G., Wattam, A. R. 1998. An important step forward in
genetic manipulation of mosquito vectors of human disease. Proceedings of the National Academy of Sciences 95:3349 – 3350.
Kline, T. C., Goering, J. J., Mathisen, O. A., Poe, P. H. 1990. Recycling
of elements transported upstream by runs of pacific salmon: 1.
15
N and 13C evidence in Sashin Creek, southeastern Alaska.
Canadian Journal of Fisheries and Aquatic Sciences 47:136 – 144.
Kohler, S. L. 1985. Identification of stream drift mechanisms: an experimental and observational approach. Ecology 66:1749 – 1761.
Lamberti, G. A. 1993. Grazing experiments in artificial streams, in
Lamberti, G. A., Steinman, A. D., Eds., Research in artificial
streams: applications, uses, and abuses. Journal of the North
American Benthological Society 12:313 – 384, pp. 337 – 342.
Lamberti, G. A. 1996. The role of periphyton in benthic food webs,
in Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Algal
ecology: freshwater benthic ecosystems. Academic Press, San
Diego, CA, pp. 533 – 572.
Lamberti, G. A., Ashkenas, L. R., Gregory, S. V., Steinman, A. D.
1987a. Effects of three herbivores on periphyton communities
in laboratory streams. Journal of the North American Benthological Society 6:92 – 104.
Lamberti, G. A., Berg, M. B. 1995. Invertebrates and other benthic
features as indicators of environmental change in Juday Creek,
Indiana. Natural Areas Journal 15:249 – 258.
Lamberti, G. A., Feminella, J. W., Resh, V. H. 1987b. Herbivory and
intraspecific competition in a stream caddisfly population. Oecologia 73:75 – 81.
Lamberti, G. A., Gregory, S. V., Ashkenas, L. R., Wildman, R. C.,
Moore, K. M. S. 1991. Stream ecosystem recovery following a
catastrophic debris flow. Canadian Journal of Fisheries and
Aquatic Sciences 48:196 – 208.
Lamberti, G. A., Gregory, S. V., Hawkins, C. P., Wildman, R. C.,
Ashkenas, L. R., DeNicola, D. M. 1992. Plant-herbivore interactions in streams near Mount St Helens. Freshwater Biology
27:237 – 247.
Lamberti, G. A., Moore, J. W. 1984. Aquatic insects as primary consumers, in Resh, V. H., Rosenberg, D. M., Eds., The ecology of
aquatic insects. Praeger, New York, pp. 164 – 195.
Lamberti, G. A., Resh, V. H. 1979. Substrate relationships, spatial
distribution patterns, and sampling variability in a stream caddisfly population. Environmental Entomology. 8:561 – 567.
772
A. E. Hershey and G. A. Lamberti
Lamberti, G. A., Resh, V. H. 1983a. Geothermal effects on stream
benthos: separate influences of thermal and chemical components on periphyton and macroinvertebrates. Canadian Journal
of Fisheries and Aquatic Sciences 40:1995 – 2009.
Lamberti, G. A., Resh, V. H. 1983b. Stream periphyton and insect
herbivores: an experimental study of grazing by a caddisfly population. Ecology 64:1124 – 1135.
Lamberti, G. A., Resh, V. H. 1985. Distribution of benthic algae and
macroinvertebrates along a thermal stream gradient. Hydrobiologia 128:13 – 21.
Lange, W. H. 1996. Aquatic and semiaquatic Lepidoptera, in Merritt, R. W., Cummins, K. W., Eds., An introduction to the
aquatic insects of North America, 3rd ed. Kendall / Hunt,
Dubuque, IA, pp. 387 – 398.
Lenat, D. R. 1988. Water quality assessment of streams using a qualitative collection method for benthic macroinvertebrates. Journal of North American Benthological Society 7:222 – 233.
Lenat, D. R., Barbour, M. T. 1994. Using benthic macroinvertebrate
community structure for rapid, cost effective, water quality
monitoring: Rapid bioassessment, in Loeb, S. L., Spacie, A.,
Eds., Biological monitoring of aquatic systems. Lewis Pubs.,
Boca Raton, FL, pp. 187 – 215.
Li, J. L., Gregory, S. V. 1989. Behavioral changes in the herbivorous
caddisfly Dicosmoecus gilvipes (Limnephilidae). Journal of the
North American Benthological Society 8:250 – 259.
Lock, M. A., Wallace, R. R., Costerton, J. W., Ventullo, R. M.,
Charlton, S. E. 1984. River epilithon: toward a structural-functional model. Oikos 42:10 – 22.
Lodge, D. M. 1991. Herbivory on freshwater macrophytes. Aquatic
Botany 41:195 – 224.
Loeb, S. L. 1994. An ecological context for biological monitoring, in
Loeb, S. L., Spacie, A., Eds., Biological monitoring of aquatic
systems. Lewis Pubs., Boca Raton, FL, pp. 3 – 7.
Lutz, P. E. 1974. Effects of temperature and photoperiod on larval
development in Tetragoneurai cynosura (Odonata: Libellulidae). Ecology 55:370 – 377.
Madsen, B. L., Bengtson, J., Butz, I. 1973. Observations on upstream
migration by imagines of some Plecoptera and Ephemeroptera.
Limnology and Oceanography 18:678 – 681.
Malmqvist, B. 1993. Interactions in stream leaf packs: effects of a
stonefly predator on detritivores and organic matter processing.
Oikos 66:454 – 462.
Mathisen, O. A., Parker, P. L., Goering, J. J., Kline, T. C., Poe, P. H.,
Scalan, R. S. 1988. Recycling of marine elements transported
into freshwater by anadromous salmon. Verhandlungen der Internationalen Vereinigung für Theoretische und Angewandte
Limnologie 23:2249 – 2258.
Mayer, M. S., Likens, G. E. 1987. The importance of algae in a
shaded headwater stream as food for an abundant caddisfly
(Trichoptera). Journal of the North American Benthological Society 6:262 – 269.
McAuliffe, J. R. 1984. Competition for space, disturbance, and the
structure of a benthic stream community. Ecology 65:894 – 908.
McCafferty, W. P. 1981. Aquatic entomology: the fisherman’s and
ecologist’s illustrated guide to insects and their relatives, Jones
and Bartlett, Boston, MA, 447 pp.
McIntire, C. D. 1973. Periphyton dynamics in laboratory streams: a
simulation model and its implications. Ecological Monographs
43:399 – 420.
McIntire, C. D., Colby, J. A. 1978. A hierarchical model of lotic
ecosystems. Ecological Monographs 48:167 – 190.
McPeek, M. A., Peckarsky, B. L. 1998. Life histories and the
strengths of species interactions: combining mortality, growth,
and fecundity effects. Ecology 79:867 – 879.
McPeek, M. A., Schrot, A. K., Brown, J. M. 1996. Adaptation to
predators in a new community: swimming performance and
predator avoidance in damselflies. Ecology 77:617 – 629.
Merritt, R. W., Cummins, K. W. 1996a. Trophic relations of
macroinvertebrates, in Hauer, F. R., Lamberti, G. A., Eds.,
Methods in stream ecology. Academic Press, San Diego, CA, pp.
453 – 474.
Merritt, R. W., Cummins, K. W., Eds. 1996b. An introduction to the
aquatic insects of North America, 3rd ed., Kendall / Hunt,
Dubuque, IA, 862 p.
Merritt, R. W., Dadd, R. H., Walker, E. D. 1992. Feeding behavior,
natural food, and nutritional relationships of larval mosquitoes.
Annual Review of Entomology 37:349 – 376.
Merritt, R. W., Lawson, D. L. 1992. The role of macroinvertebrates
in stream – floodplain dynamics. Hydrobiologia 248:65 – 77.
Meyer, J. L. 1990. A blackwater perspective on riverine ecosystems.
BioScience 40:643 – 651.
Mihuc, T. B. 1997. The functional trophic role of lotic primary consumers: generalist versus specialist strategies. Freshwater Biology 37:455 – 462.
Miller, M. C., Hershey, A. E., Merritt, R. W. 1998. Feeding behavior
of larval black flies and FPOM retention in a high gradient
blackwater stream. Canadian Journal of Zoology 76:228 – 235.
Minagawa, M., Wada, E. 1984. Stepwise enrichment of 15N along
food chains: further evidence and the relation between 15N and
animal age. Geochimica et Cosmochimica Acta 48:1135 – 1140.
Minshall, G. W. 1978. Autotrophy in stream ecosystems. Bioscience
28:767 – 771.
Minshall, G. W., Cummins, K. W., Petersen, R. C., Cushing, C. E.,
Bruns, D. A., Sedell, J. R., Vannote, R. L. 1985. Developments
in stream ecosystem theory. Canadian Journal of Fisheries and
Aquatic Sciences 42:1045 – 1055.
Minshall, G. W., Petersen, R. C., Cummins, K. W., Bott, T. L., Sedell,
J. R., Cushing, C. E., Vannote, R. L. 1983. Interbiome comparison of stream ecosystem dynamics. Ecological Monographs
53:1 – 25.
Mittelbach, G. G. 1988. Competition among refuging sunfishes and
effects of fish density on littoral zone invertebrates. Ecology
69:614 – 623.
Morse, J. C., Chapin, J. W., Herlong, D. D., Harvey, R. S. 1983.
Aquatic insects of Upper Three Runs Creek, Savannah River
Plant, South Carolina, Part II: Diptera. Journal of the Georgia
Entomological Society 18:303 – 316.
Müller, K. 1954. Investigations on the organic drift in North Swedish
streams. Institute of Freshwater Research, Drottingholm. Report No. 34:133 – 148.
Naiman, R. J., Anderson, E. C. 1996. Streams and rivers of the
coastal temperate rain forest of North America: Physical and biological variability, in Schoomaker, P. K., von Hagen, B., Eds.,
The rain forests of home: an exploration of people and place.
Island Press, Washington, DC, pp. 131 – 148.
Naiman, R. J., Beechie, T. J., Benda, L. E., Berg, D. R., Bisson, P. A.,
MacDonald, L. H., O’Connor, M. D., Olson, P. L., Steel, E. A.
1993. Fundamental elements of ecologically healthy watersheds
in the Pacific Northwest coastal ecoregion, in Naiman, R. J.,
Ed., Watershed management. Springer, New York, pp.
127 – 188.
Naiman, R. J., Melillo, J. M., Lock, M. A., Ford, T. E., Reice, S. R.
1987. Longitudinal patterns of ecosystem processes and community structure in a subarctic river continuum. Ecology
68:1138 – 1156.
Newbold, J. D., Elwood, J. W., O’Neill, R. V., Sheldon, A. L. 1983.
Phosphorus dynamics in a woodland stream ecosystem: a study
of nutrient spiralling. Ecology 64:1249 – 1265.
18. Aquatic Insect Ecology
Newman, R. M. 1991. Herbivory and detritivory on freshwater
macrophytes by invertebrates: a review. Journal of North American Benthological Society 10:89 – 114.
O’Hop, J., Wallce, J. B., Haefner, J. D. 1984. Production of a stream
shredder, Peltoperla maria (Plecoptora: Peltoperlidae) in disturbed and undisturbed hardwood catchments. Freshwater Biology 14:13 – 21.
Peckarsky, B. L. 1980. Predator-prey interactions between stoneflies
and mayflies: behavioral observations. Ecology 61:932 – 943.
Peckarsky, B. L. 1982. Aquatic insect predator-prey relations. BioScience 32:261 – 266.
Peckarsky, B. L. 1996. Alternative predator avoidance syndromes of
stream-dwelling mayfly larvae. Ecology 77:1888 – 1905.
Peckarsky, B. L., Cooper, S. D., McIntosh, A. R. 1997. Extrapolating
from individual behavior to populations and communities in
stream organisms. Journal of the North American Benthological
Society 16:375 – 390.
Peckarsky, B. L., Cowan, C. A., Penton, M. A., Anderson, C. 1993.
Sublethal consequences of stream-dwelling predatory stoneflies
on mayfly growth and fecundity. Ecology 74:1836 – 1846.
Peckarsky, B. L., Dodson, S. I. 1980. Do stonefly predators influence
benthic distributions in streams? Ecology 61:1275 – 1282.
Petersen, R. C. 1986. In situ particle generation in a southern
Swedish stream. Limnology and Oceangraphy 31:432 – 437.
Peterson, B. J., Deegan, L., Helfrich, J., Hobbie, J., Hullar, M.,
Moller, B., Ford, T., Hershey, A., Hiltner, A., Kipphut, G., Lock,
M. A., Fiebig, D. M., McKinley, V., Miller, M. C., Vestal, J. R.,
Ventullo, R., Volk, G. 1993. Biological responses of a tundra
river to fertilization. Ecology 74:653 – 672.
Peterson, B. J., Fry, B. 1987. Stable isotopes in ecosystem studies. Annual Review of Ecology and Systematics 18:293 – 320.
Plafkin, J. L., Barbour, M. T., Porter, K. D., Gross, S. K., Hughes,
R. M. 1989. Rapid bioassessment protocols for use in streams
and rivers. U. S. EPA, Office of Water, EPA / 444 / 4-89-001.
Pritchard, G. 1983. Biology of Tipulidae. Annual Review of Entomology 28:1 – 22.
Pritchard, G. 1991. Insects in thermal springs. Memoirs of the Entomological Society of Canada 155:89 – 106.
Pringle, C. M. 1985. Effects of chironomid (Insecta: Diptera) tube
building activities on stream diatom communities. Journal of
Phycology 21:185 – 194.
Power, M. E. 1990. Effects of fish in river food webs. Science
250:811 – 814.
Rader, R. B., Ward, J. V. 1987. Resource utilization, overlap and
temporal dynamics in a guild of mountain stream insects. Freshwater Biology 18:521 – 528.
Ramírez, A., Pringle, C. M. 1998. Structure and production of a benthic insect assemblage in a neotropical stream. Journal of the
North American Benthological Society 17:443 – 463.
Rasmussen, J. B. 1988. Habitat requirements of burrowing mayflies
(Ephemeridae: Hexagenia) in lakes, with special reference to the
effects of eutrophication. Journal of the North American Benthological Society 7:51 – 64.
Reice, S. R. 1983. Predation and substratum: factors in lotic community structure, in Fontaine, T. F. III, Bartell, S. M., Eds., Dynamics of lotic ecosystems. Ann Arbor Science, Ann Arbor, MI, pp.
325 – 345.
Renshaw, M., Elias, M., Maheswary, N. P., Hassan, M. M., Silver, J. B.,
Birley, M. H. 1996. A survey of larval and adult mosquitoes on
the flood plains of Bangladesh, in relation to flood-control activities. Annals of Tropical Medicine and Parasitology 90:621 – 634.
Resh, V. H. 1992. Year-to-year changes in the age structure of a caddisfly population following loss and recovery of a springbrook
habitat. Ecography 15:314 – 317.
773
Resh, V. H., Brown, A. V., Covich, A. P., Gurtz, M. E., Li, H. W.,
Minshall, G. W., Reice, S. R., Sheldon, A. L., Wallace, J. B.,
Wissmar, R. 1988. The role of disturbance in stream ecology.
Journal of the North American Benthological Society 7:433 – 455.
Resh, V. H., Flynn, T. S., Lamberti, G. A., McElravy, E. P., Sorg,
K. L., Wood, J. R. 1981. Responses of the sericostomatid caddisfly Gumaga nigricula (McL.) to environmental disruption.
Series Entomologica 20:311 – 318.
Resh, V. H., Houp, R. E. 1986. Life history of the caddisfly Dibusa
angata and its association with the red alga Lemanea australis.
Journal of the North American Benthological Society 5:28 – 40.
Resh, V. H., Lamberti, G. A., Wood, J. R. 1984. Biology of the caddisfly Helicopsyche borealis (Hagen): a comparison of North American populations. Freshwater Invertebrate Biology 3:172 – 180.
Resh, V. H., Myers, M. J., Hannaford, M. J. 1996. Macroinvertebrates as biotic indicators of environmental quality, in Hauer,
F. R., Lamberti, G. A., Eds., Methods in stream ecology. Academic Press, San Diego, CA, pp. 647 – 667.
Richards, C., Host, G. E. 1994. Examining land use influences on
stream habitats and macroinvertebrates: a GIS approach. Water
Resources Bulletin 30:729 – 738.
Richards, C., Host, G. E., Arthur, J. W. 1993. Identification of predominant environmental factors structuring stream macroinvertebrate communities within a large agricultural catchment.
Freshwater Biology 29:285 – 294.
Richards, C., Johnson, L. B., Host, G. 1996. Landscape scale influences on stream habitats and biota. Canadian Journal of Fisheries and Aquatic Sciences 53:295 – 311.
Richards, C., Minshall, G. W. 1992. Spatial and temporal trends in
stream macroinvertebrate species assemblages: the influence of
watershed disturbance. Hydrobiologia 241:173 – 184.
Richardson, C. J. 1979. Primary productivity values in freshwater
wetlands, in Greeson, P. E., Clark, J. R., Clark, J. E. Eds., Wetland function and values: the state of our understanding . American Water Resources Association, Minneapolis, MN, pp.
131 – 145.
Richardson, J. S. 1993. Limits to productivity in streams: evidence
from studies of macroinvertebrates. Canadian Special Publications in Fisheries and Aquatic Sciences 118:9 – 15.
Richey, J. E., Perkins, M. A., Goldman, C. R. 1975. Effects of Kokanee salmon (Oncorhynchus nerka) decomposition on the ecology of a subalpine stream. Journal of the Fisheries Research
Board of Canada 32:817 – 820.
Rosenberg, D. M., Danks, H. V. 1987. Aquatic insects of peatlands
and marshes in Canada: introduction. Memoirs of the Entomological Society of Canada 140:1 – 4.
Rosenberg, D. M., Resh, V. H. 1996. Use of aquatic insects in biomonitoring. in Merritt, R. W., Cummins, K. W., Eds., An introduction to the aquatic insects of North America, 3rd ed.
Kendall / Hunt, Dubuque, IA, pp. 87 – 97.
Rosenberg, D. M., Wiens, A. P., Saether, O. A. 1977. Life histories of
Cricotopus (Cricotopus) bicinctus and C. (C.) mckenziensis
(Diptera: Chironomidae) in the Fort Simpson area, Northwest
Territories. Journal of the Fisheries Research Board of Canada
34:247 – 253.
Ross, D. H., Wallace, J. B. 1981. Production of Brachycentrus spinae
Ross (Trichoptera: Brachycentridae) and its role in seston dynamics of a sourthern Applacahain stream (USA). Holarctic
Ecology 6:270 – 284.
Saether, O. A. 1979. Chironomid communities as water quality indicators. Holarctic Ecology 2:65 – 74.
Saponis, A. R. 1977. A revision of the Nearctic species of Orthcladius (Orthocladius) van der Wulp (Diptera: Chironomidae).
Memoirs of the Entomological Society of Canada 102:1 – 187.
774
A. E. Hershey and G. A. Lamberti
Schindler, D. W. 1977. The evolution of phosphorus limitation in
lakes. Science 195:260 – 262.
Schmidt, S. K., Huges, J. M., Bunn, S. E. 1995. Gene flow among
conspecific populations of Baetis sp. (Ephemeroptera): adult
flight and larval drift. Journal of the North American Benthological Society 14:147 – 157.
Schuldt, J. A., Hershey, A. E. 1995. Impact of salmon carcass decomposition on Lake Superior tributary streams. Journal of the
North American Benthological Society 14:259 – 268.
Scrimgeour, G. J., Culp, J. M. 1994. Feeding while evading predators
by a lotic mayfly: linking short-term foraging behavior to longterm fitness consequences. Oecologia 199:128 – 134.
Sedell, J. R., Reeves, G. H., Hauer, F. R., Standford, J. A., Hawkins,
C. P. 1990. Role of refugia in recovery from disturbances: modern fragmentation and disconnected river systems. Environmental Management 14:711 – 724.
Shepard, R. B., Minshall, G. W. 1981. Nutritional value of lotic insect feces compared with allocthonous materials. Archiv fur Hydrobiologie 90:467 – 488.
Smock, L. A. 1996. Macroinvertebrate movements: drift, colonization, and emergence, in Hauer, F. R., Lamberti, G. A., Eds.,
Methods in Stream Ecology. Academic Press, San Diego, CA,
pp. 371 – 390.
Smock, L. A., Roeding, C. E. 1986. The trophic basis of production
of the macroinvertebrate community of a southeastern U.S.A.
blackwater stream. Holarctic Ecology 9:165 – 174.
Smock, L. A., Gladden, J. E., Riekenberg, J. L., Smith, L. C., Black,
C. R. 1992. Lotic macroinvertebrate production in three dimensions: channel surface, hyporheic, and floodplain environments.
Ecology 73:876 – 886.
Soluk, D. A. 1985. Macroinvertebrate abundance and production of
psammophilous Chironomidae in shifting sand areas of a lowland river. Canadian Journal of Fisheries and Aquatic Sciences
42:1296 – 1302.
Sousa, W. P. 1984. The role of disturbance in natural communities.
Annual Review of Ecology and Systematics 15:353 – 391.
Spence, J. R., Andersen, N. M. 1994. Biology of water striders: interactions between systematics and ecology. Annual Review of
Entomology 39:101 – 128.
Stanford, J. A., Ward, J. V. 1988. The hyporheic habitat of river
ecosystems. Nature 335:64 – 66.
Steinman, A. D. 1996. Effects of grazers on freshwater benthic algae,
in Stevenson, R. J., Bothwell, M. L., Lowe, R. L., Eds., Algal
ecology: freshwater benthic ecosystems. Academic Press, San
Diego, CA, pp. 341 – 373.
Stewart, A. J., Loar, J. M. 1994. Spatial and temporal variation in
biological monitoring data, in Loeb, S. L., Spacie, A., Eds.,
Biological Monitoring of Aquatic Systems. Lewis Publishers,
Boca Raton, FL, pp. 91 – 124.
Stewart, K. W., Harper, P. P. 1996. Plecoptera, in Merritt, R. W.,
Cummins, K. W., Eds., An introduction to the aquatic insects of
North America, 3rd ed. Kendall / Hunt, Dubuque, IA, pp.
217 – 266
Stockner, J. G., Shortreed, K. R. S. 1978. Enhancement of autotrophic production by nutrient addition in a coastal rainforest
stream on Vancouver Island. Journal of the Fisheries Research
Board of Canada 35:28 – 34.
Sullivan, K., Lisle, T. E., Dolloff, C. A., Grant, G. E., Reid, L. M.
1987. Stream channels: the link between forests and fishes, in
Salo, E. O., Cundy, T. W., Eds., Streamside management:
forestry and fishery interactions. University of Washington Institute of Forest Resources Contribution No. 57, pp. 39 – 97.
Suren, A. M., Winterbourn, M. J. 1992. The influence of periphyton,
detritus and shelter on invertebrate colonization of aquatic
bryophytes. Freshwater Biology 27:327 – 339.
Swanson, F. J., Benda, L. E., Duncan, S. H., Grant, G. E., Megahan,
W. F., Reid, L. M., Zeimer, R. R. 1987. Mass failures and other
processes of sediment production in Pacific Northwest forest
landscapes, in Salo, E. O., Cundy, T. W., Eds., Streamside management: forestry and fishery interactions. University of Washington Institute of Forest Resources Contribution No. 57,
pp. 9 – 38.
Swanson, F. J., Gregory, S. V., Sedell, J. R., Campbell, A. G. 1982.
Land – water interactions: the riparian zone, in Edmonds, R. L.,
Ed., Analysis of coniferous forest ecosystems in the western
United States. US / IBP Synthesis Series 14. Hutchinson-Ross,
Stroudsburg, PA, pp. 267 – 291.
Swanson, G. A. 1984 / 1985. Invertebrates consumed by dabbling
ducks (Anatinae) on the breeding grounds. Jounnal of the Minnesota Academy of Science 50:37 – 40.
ter Braak, C. J. F., Prentice, I. C. 1988. A theory of gradient analysis.
Advances in Ecological Research 18:271 – 317.
Tester, J. 1995. Minnesota’s natural heritage: an ecological perspective. Univ. of Minnesota Press. Minneapolis, MN.
Thorp, J. H. 1986. Two distinct roles for predators in freshwater
assemblages. Oikos 47:75 – 82.
Thorp, J. H., Bergey, E. A. 1981. Field experiments on responses of a
freshwater, benthic macroinvertebrate community to vertebrate
predators. Ecology 62:365 – 375.
Thorp, J. H., Delong, M. D. 1994. The riverine productivity model:
an heuristic view of carbon sources and organic processing in
large river ecosystems. Oikos 70:305 – 308.
Thorp, J. H., Diggins, M. R. 1982. Factors affecting depth distribution of dragonflies and other benhtic insects in a thermally
destabilized reservoir. Hydrobiologia 87:33 – 44.
Ulfstrand, S. 1968. Benthic animals in Lapland streams. Oikos
Supplement 10:1 – 120.
Usinger, R. L. 1956. Aquatic insects of California, Univ. of California
Press, Berkeley, CA, 508 p.
Van Buskirk, J. 1989. Density-dependent cannibalism, and size
class dominance in a dragonfly Aeshna juncea. Ecology 74:
1950 – 1958.
Vander Zanden, M. J., Rasmussen, J. B. 1999. Primary consumer
13C and 15N and the trophic position of aquatic consumers.
Ecology 80:1242 – 1252.
Vannote, R. L., Minshall, G. W., Cummins, K. W., Sedell, J. R.,
Cushing, C. E. 1980. The river continuum concept. Canadian
Journal of Fisheries and Aquatic Sciences 37:130 – 137.
Van Sickle, J., Gregory, S. V. 1990. Modeling inputs of coarse woody
debris to streams from falling trees. Canadian Journal of Forest
Research 20:1593 – 1601.
Vodopich, D. S., Cowell, B. C. 1984. Interaction of factors controlling the distribution of a predatory insect. Ecology 65:39 – 52.
Walker, E. D., Merritt, R. W. 1988. The significance of leaf detritus
to mosquito (Diptera: Culicidae) productivity from treeholes.
Environmental Entomology 17:199 – 206.
Wallace, J. B., Brady, U. E. 1971. Residue levels of dieldrin in aquatic
invertebrates and effect of prolonged exposure on populations.
Pesticides Monitoring Journal 5:295 – 300.
Wallace, J. B., Merritt, R. W. 1980. Filter-feeding ecology of aquatic
insects. Annual Review of Entomology 25:103 – 132.
Wallace, J. B., Webster, J. R., Cuffney, T. F. 1982. Stream detritus
dynamics: regulation by invertebrate consumers. Oecologia
53:197 – 200.
Wallace, J. B., O’Hop, J. 1985. Life on a fast pad: waterlily leaf beetle impact on water lilies. Ecology 66:1534 – 1544.
Wallace, J. B., Vogel, D. S., Cuffney, T. F. 1986. Recovery of a headwater stream from an insecticide-induced community disturbance. Journal of the North American Benthological Society
5:115 – 126.
18. Aquatic Insect Ecology
Wallace, J. B., Gurtz, M. E., Smith-Cuffney, F. 1988. Long-term comparisons of insect abundances in disturbed and undisturbed
Appalachian headwater streams. Verhandlungen Internationale
Verein Limnologie 23:1224 – 1231.
Wallace, J. B., Lugthart, G. J., Cuffney, T. F., Schurr, G. A. 1989. The
impact of repeated insecticide treatments on drift and benthos
of a headwater stream. Hydrobiologia 179:135 – 147.
Wallace, J. B., Anderson, N. H. 1996. Habitat, life history, and behavioral adaptations of aquatic insects, in Merritt, R. W., Cummins,
K. W., Eds., An Introduction to the Aquatic Insects of North
America, 3rd ed. Kendall / Hunt, Dubuque, IA, pp. 41 – 73.
Wallace, J. B., Grubaugh, J. W. 1996. Transport and storage of
FPOM, in Hauer, F. R., Lamberti, G. A., Eds., Methods in
stream ecology, Academic Press, San Diego, CA, pp. 191 – 215.
Wallace, J. B., Webster, J. R. 1996. The role of macroinvertebrates in
stream ecosystem function. Annual Review of Entomology
41:115 – 139.
Wallace, J. B., Eggert, S. L., Meyer, J. L., Webster, J. R. 1997. Multiple trophic levels of a forest stream linked to terrestrial litter inputs. Science 277:102 – 104.
Walshe, B. M. 1947. Feeding mechanisms of Chironomus larvae. Nature 160:474.
Ward, A. K., Dahm, C. N., Cummins, K. W. 1985. Nostoc
(Cyanophyta) productivity in Oregon stream ecosystems: invertebrate influences and differences between morphological types.
Journal of Phycology 21:223 – 227.
Ward, G. M., Ward, A. K., Dahm, C. N., Aumen, N. G. 1994. Origin
and formation of organic and inorganic particles in aquatic systems, in Wotton, R. S., Ed., The biology of particles in aquatic
systems, 2nd ed. Lewis Pubs., Boca Raton, FL, pp. 27 – 56.
Ward, J. V. 1992. Aquatic insect ecology. 1. Biology and habitat.
Wiley, New York, NY.
Ward, J. V., Stanford, J. A. 1982. Thermal responses in the evolutionary ecology of aquatic insects. Annual Review of Entomology 27:97 – 117.
Ward, J. V., Stanford, J. A. 1983. The intermediate disturbance hypothesis: an explanation for biotic diversity patterns in lotic ecosystems, in Fontaine, T. D., Bartell, S. M., Eds., Dynamics of lotic
ecosystems. Ann Arbor Science, Ann Arbor, MI, pp. 347 – 356.
Warwick, W. F. 1980. Paleobiology of the Bay of Quinte, Lake
Ontario: 2800 years of cultural influence. Canadian Bulletin of
Fisheries and Aquatic Sciences 206:1 – 117.
Warwick, W. F., Tisdale, N. A. 1989. Morphological deformities
in Chironomus, Cryptochironomus, and Procladius larvae
(Diptera: Chironomidae) from two differentially stressed sites in
Tobin Lake, Saskatchewan. Canadian Journal of Fisheries and
Aquatic Sciences 45:1123 – 1144.
Waters, T. F. 1972. The drift of stream insects. Annual Review of
Entomology 17:253 – 272.
Waters, T. F. 1977. Secondary production in inland waters. Advances
in Ecological Research 10:91 – 164.
Waters, T. F., Crawford, G. W. 1973. Annual production of a stream
mayfly population: A comparison of methods. Limnology and
Oceanography 18:286 – 296.
775
Waters, T. F., Hokenstrom, J. C. 1980. Annual production and drift
of the stream amphipod Gammarus pseudolimnaeus in Valley
Creek, Minnesota. Limnology and Oceanography 25:700 – 710.
Wellborn, G. A., Skelly, D. K., Werner, E. E. 1996. Mechanisms creating community structure across a freshwater habitat gradient.
Annual Review of Ecology and Systematics 27:337 – 367.
Wheeler, J. R. 1994. Factors affecting black fly abundance and distribution in an arctic stream. M. S. Thesis, University of Minnesota, Duluth, MN.
Whiles, M. R., Wallace, J. B., Chung, K. 1993. The influence of Lepidostoma (Trichoptera: Lepidostomatidae) on recovery of leaflitter processing in disturbed headwater streams. The American
Midland Naturalist 130:356 – 363.
Whitledge, G. W., Rabeni, C. K. 1997. Energy sources and ecological
role of crayfishes in an Ozark stream: insights from stable isotopes and gut analyses. Canadian Journal of Fisheries and
Aquatic Sciences 54:2555 – 2563.
White, D. S., Brigham, W. U. 1996. Aquatic Coleoptera, in Merritt,
R. W., Cummins, K. W., Eds., An introduction to the aquatic
insects of North America, 3rd ed., Kendall / Hunt, Dubuque, IA,
pp. 399 – 473.
Wiggins, G. B. 1996. Larvae of the North American caddisfly genera
(Trichoptera), 2nd ed., Univ. of Toronto Press, Toronto, 457 p.
Wiggins, G. B., Mackay, R. J., Smith, I. M. 1980. Evolutionary and
ecological strategies of animals in temporary pools. Archiv fur
Hydrobiologie / Supplementum 58:97 – 206.
Williams, D. D. 1993. Nutrient and flow vector dynamics at the
hyporheic / groundwater interface and their effects on the interstitial fauna. Hydrobiologia 251:185 – 198.
Williams, D. D., Feltmate, B. W. 1992. Aquatic insects. C.A.B. Int.,
Wallingford, Oxon, UK.
Wilzbach, M. A., Cummins, K. W. 1989. An assessment of shortterm depletion of stream macroinvertebrate benthos by drift.
Hydrobiologia 185:29 – 39.
Winterbourn, M. J., Rounick, J. S., Cowie, B. 1981. Are New
Zealand stream really different? New Zealand Journal of
Marine and Freshwater Research 15:321 – 328.
Wissinger, S. A. 1988. Life history and size structure of larval dragonfly populations. Journal of the North American Benthological
Society 7:13 – 28.
Wolheim, W. M., Peterson, B. J., Deegan, L. A., Bahr, M., Jones, D.,
Bowden, W. B., Hershey, A. E., Kling, G. W., Miller, M. C.
1999. A coupled field and modeling approach for the analysis
of nitrogen cycling in streams. Journal of the North American
Benthological Society 18:199 – 221.
Wooster, D. 1994. Predator impacts on stream benthic prey. Oecologia 99:7 – 15.
Wotton, R. S. 1976. Evidence that blackfly larvae can feed on particles of colloidal size. Nature 261:697.
Wotton, R. S. 1988. Dissolved organic material and trophic dynamics. BioScience 38:172 – 177.
Zachariassen, K. E. 1979. The mechanism of the cryoprotective effect of glyceral in beetles tolerant to freezing. J. Insect Physiol.
25:29 – 32.
This Page Intentionally Left Blank
19
INTRODUCTION TO THE
SUBPHYLUM CRUSTACEA
Alan P. Covich
James H. Thorp
Department of Fishery and Wildlife Biology
Colorado State University
Fort Collins, Colorado 80523
Department of Biology
Clarkson University
Potsdam, New York 13699
I. Introduction
II. Anatomy and Physiology of Crustacea
A. External Morphology
B. Organ System Function
C. Environmental Physiology
III. Ecology and Evolution of
Selected Crustacea
A. General Relationships
B. Ecological Distributions
and Interactions
C. Life-History Traits
D. Biogeography
IV. Collecting, Rearing, and Preparation
for Identification
V. Classification of Peracarida and
Brachiura
A. Taxonomic Key
Acknowledgments
Literature Cited
I. INTRODUCTION
Crustaceans are the most diverse of any group of
arthropods and currently thrive in a wide array of habitats. Most species occur primarily in aquatic habitats but
some are adapted to live on land. The fossil record extends back to the lower Cambrian and throughout most
of this long period, crustacean fossils are associated with
a broad range of aquatic habitats, especially marine waters (Schram, 1982). Groups of crustaceans adapted to
freshwater habitats very early in their evolutionary history (Hutchinson, 1967; Abele, 1982). The fossil record
for freshwater species is especially well preserved from
the Tertiary for some groups that are clearly related to
living forms. Although only about 10% of the nearly
40,000 extant species in the subphylum Crustacea (phylum Arthropoda) occur in the inland pools, lakes, and
streams of the world (Table I) (Bowman and Abele,
1982; Banarescu, 1990; Barnes and Harrison, 1992),
freshwater crustaceans are extremely important in many
ecosystem processes of inland waters.
Crustaceans have evolved into several major
groups (Fig. 1) that represent distinct modifications for
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
exploiting resources and performing ecosystem services
in many different surface waters and subsurface habitats. Amphipods and isopods, as well as mysids and
branchiura are discussed in this chapter; ostracodes are
reviewed in Chapter 20. The cladocerans and copepods
together constitute the dominant component of most
freshwater planktonic assemblages and are discussed in
Chapters 21 and 22 respectively.
Crustacean zooplankton are major consumers of
phytoplankton and they transfer energy from these primary producers up the food web to zooplanktivorous
fish in lakes, ponds, and rivers. Some crustacean zooplankton also consume aggregations formed from dissolved organic matter and other organic particulates.
Others are predatory on rotifers, protozoans, other
crustaceans, aquatic insects, and fish eggs and larvae.
Some zooplankton vertically migrate and thereby link
deep, benthic habitats with open waters of lakes. These
migratory species are important in nutrient cycling and
transfers of toxins (such as heavy metals and pesticides)
from sediments to pelagic foodwebs. Along with many
other invertebrates, crustaceans provide hard surfaces
as semipermanent, mobile substrates for diverse
777
778
A. P. Covich and J. H. Thorp
TABLE I Classification of Crustacea and Estimates of Species Numbers in North American
Fresh Watersa,b
Taxa
Subphylum Crustacea
Class Cephalocarida
Class Branchiopoda
[eight orders, including cladocerans]
Class Remipdia
Class Maxillopoda
subclass Branchiura
subclass Copepoda
Class Ostracoda
Class Malacostraca
Superorder Pancarida
[order Thermosbaenacea]
Superorder Peracarida
[orders Mysidacea, Amphipoda, Isopoda]
Superorder Eucarida
[order Decapoda]
a
b
Worldwide
percentage found
in Freshwaters
Approximate
number of species
in North America
Chapter(s) of
this book
10%
0%
95%
ca.1500
0
200
19 – 23
—
21
0%
15%
90%
15%
50%
10%
50%
0
23
200
420
1
—
19, 22
19
22
20
19, 23
19
10%
300
19
334
23
10%
North of Mexico.
Classification based in part on Bowman and Abele (1982).
epibiont species such as ciliated protozoa and sessile
rotifers (chapters 3 and 8; Threlkeld et al., 1993).
Within inland hypersaline environments, brine shrimp
(Chapter 21) are sometimes the only metazoan capable
of surviving hyperosmotic conditions. As salinity fluctuates, the crustacean community can rapidly change its
species composition and exploit different sources of energy (Galat et al., 1988; Hammer et al., 1990; Stephens,
1990; Wurtsbaugh and Berry, 1990; Williams, 1998).
Crustaceans also comprise a significant portion of
the benthic biomass in many lakes and streams. Some
species are abundant in the littoral zone and throughout
the oxygenated deeper water where they feed on bacteria and fungi on decaying vegetation or live algae (Gee,
1988; Delong et al., 1993). As with the zooplankton,
these benthic consumers recycle nutrients that sustain
high levels of microbial productivity. Some species horizontally migrate at night and link the littoral zone with
pelagic water. This nocturnal mobility, like vertical migrations, can limit vulnerability of these species to visual predators. In streams, crustaceans also occupy
many different habitats ranging from deep, subsurface
waters to quiet pools and fast-flowing riffles where they
actively consume a wide range of foods (Marchant and
Hynes, 1981a, b; Boulton et al., 1998). In caves, unique
species of crustaceans have evolved that also feed on
many types of resources (Culver et al., 1995; Carpenter,
1999). Similarly, some species are found in wetlands
and these feed on many types of detrital foods (Smock
and Harlowe, 1983). These diverse benthic species
function as herbivores, carnivores, and detritivores
while also providing many ecosystem services (Palmer
et al., 1997; Boulton et al., 1998; Covich et al., 1999).
As discussed below, some of these very productive benthic species live in extreme habitats, from cold arctic
waters to hot springs and saline waters where they can
dominate benthic communities and reach high densities.
One of the most ecologically and taxonomically diverse freshwater crustacean groups north of the Rio
Grande are omnivorous crayfish (Decapoda; Chapter
23). Of the more than 500 species and subspecies currently known worldwide, 393 taxa (78%) occur in
North America and 346 of these occur in the United
States and Canada (Sket, 1999a; Taylor, 2000; see
Hobbs, Chapter 23). River drainages in the southeastern United States have the highest diversity and highest
concentrations of crayfish species of any region anywhere (Taylor, 2000). These diverse benthic consumers
are not only important in nutrient cycling, but they
also can change the life histories of some of their prey
species that have evolved ways to avoid being eaten
(Crowl and Covich, 1990). Crayfish maintain very high
densities in many different habitats where they feed on
a wide range of living and dead plants and animals. In
addition to their crucial roles as consumers and secondary producers in natural aquatic ecosystems, some
crustaceans are increasing in importance as a food
source for humans. Crayfish have become widely
available through aquacultural production, as have the
larger “river shrimp” (Macrobrachium) in warm
19. Introduction to the Subphylum Crustacea
779
FIGURE 1
General evolutionary linkages of major groups of freshwater crustaceans. From Bowman
and Abele (1982) who stated: “This [classification] is not to indicate phylogenetic relationships and
should not be so interpreted. The dashed line at the base emphasizes uncertainties concerning the origins of the five classes and their relationships to each other.”
temperate-zone and tropical locations. Benthic amphipods, isopods, ostracodes, and various littoral
cladocerans and copepods consume a large amount of
living organisms and particulate organic matter. All
these crustaceans, in turn, can be significant constituents in the diets of many invertebrate predators
and bottom-feeding fishes (Dahl, 1998; Wooster, 1998;
MacNeil et al., 1999a, b).
The present chapter provides a brief introduction to
crustacean anatomy and physiology. More detailed information can be obtained from general invertebrate
zoology texts (e.g., Barnes, 1987 and from several
books on crustaceans, such as those by Schmitt (1965),
Schram (1986), Fitzpatrick (1983), McLaughlin (1980),
and the 10-volume set edited by Bliss (1982 – 1985)
and several co-editors. Following an introduction to
780
A. P. Covich and J. H. Thorp
anatomy and physiology (based primarily on information from these sources), this chapter summarizes the
ecology and evolution of three orders of crustaceans:
Mysidacea, Amphipoda, and Isopoda. Brief reviews of
the subclass Branchiura and the class Remipedia are
also included. This chapter concludes with a discussion
of crustacean biogeography, classification, and a key to
the genera of the major taxa listed above. Related information is presented on other major crustacean groups
in Chapters 20, 21, 22, and 23.
II. ANATOMY AND PHYSIOLOGY OF CRUSTACEA
A. External Morphology
Although many other arthropod groups use a single basic body plan, the crustaceans are anatomically
very diverse (Schram, 1986). Once you have seen one
insect or spider taxon, you have a good idea of the general body form. The crustaceans, however, have
evolved a high diversity of body forms by fusing various segments or by developing highly specialized body
segments and appendages. Crustaceans are basically
metameric, protostomate coelomates whose bodies are
covered by a hard exoskeleton and often divided into
three regions, or tagmata: the head (cephalic area), thorax, and abdomen (sometimes the first two regions are
combined as a cephalothorax). The jointed appendages
are primitively biramous and may be present in all
three body regions. Mandibles, two pairs of maxillae,
and two pairs of antennae are present in all species
during at least one life stage. Crustaceans are distinguished from the Insecta, one of the other two large
arthropod groups, by the presence of two pairs of
antennae.
1. Skeleton
All crustaceans have a hard exoskeleton composed
of chitin that varies in rigidity at various life-history
stages. The name Crustacea is derived from the Latin
word for shell and refers to this exoskeleton, which in
many taxa is hardened with very small inclusions of
calcium carbonate. Although not a true “shell” as is
found among the Mollusca, this exoskeleton is very
durable. Crustaceans then, like other arthropods, are
characterized by a chitinized exoskeleton consisting of
a thin proteinaceous stiffened epicuticle and a thick,
multilayer procuticle strengthened by calcium carbonate. The skeleton attains its maximum thickness and
rigidity in the decapods. Projecting inward are chitinized struts, known as apodemes, which serve as sites
for internal attachment of the muscles; they also lend
partial protection to some organs.
2. Appendages
Crustacean appendages are modified among
species to serve a large variety of purposes, including
locomotion (walking and swimming), feeding, grooming, respiration, sensory reception, reproduction, and
defense. Consequently, the primitive, generally biramous appendages (terminal exopod and endopod) are
often extensively modified with additional lateral and
medial projections. Two extreme forms are recognized
among adults (Fig. 2): the leaflike or lobed phyllopod
appendage (as found among branchiopods) and the unbranched, segmented walking leg, or stenopod (typical
of crayfish). The cephalic region contains five basic
types of paired appendages: (1) antennules (first antennae), which are uniramous in all crustaceans except the
malacostracans; (2) antennae (second antennae); (3)
mandibles; (4) maxillules (first maxillae); and (5) maxillae (second maxillae). The first two pairs generally
have a sensory function (aiding some taxa in food location and filtering), whereas the last three pairs normally
function in food acquisition, handling, or processing.
The number of appendages on the thorax and abdomen vary greatly among large taxonomic groups.
Malacostracans (such as decapods and amphipods)
generally possess 5 – 8 pairs of thoracic appendages
(sometimes called pereiopods) and six pairs of abdominal appendages (pleopods and terminal uropods). Primary abdominal appendages are absent in all nonmalacostracans except Notostraca, although comparable
structures may have secondarily evolved.
B. Organ System Function
1. Nutrition and Digestion
Freshwater crustaceans predominantly employ their
antennae, maxillae, or thoracic appendages to filter or
grab algae, bacteria, microzooplankton, or suspended
dead organic matter (detrital seston) from the water column. In some habitats, a relatively large number of taxa
are primarily carnivores or scavengers (e.g., benthic decapods, certain copepods, and others). Parasitism is rare
except among specialized copepods and branchiurans
(Yamaguti, 1963; Margolis and Kabata, 1988) that attack fish and bopyrid isopods, such as Probopyrus which
attack freshwater shrimps (Beck, 1980; Collart, 1990;
Roman Contreras, 1996). Only one ostracod genus, Entocythere, is known to be parasitic. Cannibalism generally occurs when newly molted individuals are vulnerable, especially among the decapods (Goddard, 1988),
amphipods (Dick, 1995; MacNeil et al., 1997, 1999),
and isopods (Jormalainen and Shuster, 1997).
Using their cephalic or thoracic appendages, crustaceans acquire food and pass it through a ventral
19. Introduction to the Subphylum Crustacea
781
FIGURE 2
Representative crustacean appendanges: (A) a phyllopod appendage of Anostraca; (B) biramous
appendage of Anaspidacea (superorder Syncarida); and (C) uniramous stenopod appendage of the Stenopodidea (Decapoda: Pleocyemata). [Redrawn from McLaughlin, 1982].
mouth into a tripartite alimentary tract (see Chapter
23, Fig. 4). Crayfish have sharp mandibles that assist
in cutting and shredding their food. Crayfish chelae
(“claws”) can be used to grasp larger food items
(although they are often used for defense) and their numerous sensilla (sensory hairs) may aid in chemically
locating food (Fig. 3). In most crustaceans, food is
ground into small, digestible particles by the gastric
mill (the posterior end of the cardiac stomach) of the
muscular foregut (e.g., Schmitz and Sherrey, 1983). Enzymes (primarily from the hepatopancreas) empty into
the short midgut, where food is digested and assimilated. Wastes are conveyed by peristalsis through the
long hindgut and out the terminal anus.
2. Circulation and Respiration
Crustaceans generally have an open circulatory
system consisting of a dorsal, neurogenic heart with a
single chamber, one or more arteries (in malacostracans
only), and several sinuses for returning blood to the
heart. Many noncalanoid copepods (and marine barnacles) lack a heart altogether; fluids are circulated as a
result of general body movements. The heart is usually
located in the thorax or cephalothorax, but is present
in the abdomen in species with abdominal gills (such as
isopods). The blood contains low amounts of a vis-
cous, copper-based respiratory pigment (hemocyanin),
which is dissolved in the hemolymph.
Respiration takes place entirely across the body
integument in nonmalacostracans, but thoracic and
abdominal gills are typical of other freshwater crustaceans. Peracaridans (discussed later in this chapter)
feature rudimentary epipodal gills (flattened outgrowths of the coxa). Individual species differ in their
rates of oxygen consumption and their ability to regulate their respiration as dissolved oxygen concentrations decrease (Toman and Dall, 1998). Freshwater decapods are distinct in having trichobranchiate gills
composed of a central axis (with afferent and efferent
blood vessels) bearing numerous filamentous branches
(Fig. 4).
3. Fluid and Solute Balance
Paired maxillary and antennal glands (also called
green glands) are the principal excretory organs in
crustaceans. The “labyrinth” of the antennal gland is
also involved in reabsorption of glucose, amino acids,
and divalent ions from tubule fluids. Maxillary glands
predominate within lower crustaceans, whereas the
slightly more complex antennal glands characterize
most malacostracans. More than 90% of the nitrogenous wastes are voided as ammonia.
782
A. P. Covich and J. H. Thorp
FIGURE 3
Scanning electron micrographs of crayfish chela: (A) view of distal portion of chela
(Orconectes propinquus, female, 300 magnification); (B) Plumose sensillia (feathered hairs) located on
the cutting edge of chela (Orconectes virilis, female, 6000 magnification). [Images provided by J. L.
Borash, 1997. See Borash and Moore, 1997 for more information.]
19. Introduction to the Subphylum Crustacea
783
living in inland salt lakes. The brine shrimp Artemia
salina, tolerates salinities ranging from 10% seawater
to crystallizing brine; it is hypertonic to the medium in
dilute waters, but hypotonic in concentrated solutions.
4. Neural Systems
FIGURE 4 Structure of decapod gill branches, as shown in transverse
(upper) and latteral (lower) views: (A) dendrobranchiate; (B)
phyllobranchiate; and (C) trichobranchiate. [From McLaughlin, 1982].
Freshwater crustaceans are relatively competent
osmoregulators. They face problems of water influx
and salt loss (both primarily across the gills) and, therefore, must release copious quantities of urine that is
isosmotic to the hemolymph (a few taxa produce urine
that is hyposmotic to the blood). Freshwater branchiopods and some copepods maintain a dilute hemolymph equal to about 200 mOsm (20% of seawater), while the blood of crayfish is more salty (about
350 mOsm; see Mantel and Farmer, 1983). An opposite situation is faced by hyporegulating crustaceans
FIGURE 5
The crustacean neural system is comprised of a tripartite brain and paired ventral, ganglionated nerve
cords linked by commissures in a ladderlike arrangement. The protocerebrum of the brain normally innervates the eye, sinus gland, frontal organs, and head muscles; the deutocerebrum controls the antennules; the
tritocerebrum innervates the antennae and a portion of
the alimentary tract. Crustaceans are generally sensitive
to light, chemicals, temperature, touch, gravity, pressure,
and sound. The eyes of adult malacostracans are compound and frequently mounted on a stalk, or peduncle;
but the adult eye of many entomostracans (an older
term inclusive of most nonmalacostracans) is not much
advanced over the simple cluster of inverse pigment cup
ocelli that characterizes the larval (naupliar) eye.
5. Reproduction, Development, and Growth
Crustaceans are primarily sexually reproducing,
dioecious organisms, but hermaphroditism and parthenogenesis occur sporadically among entomostracans (especially in ostracodes and branchiopods) and a few
marine malacostracans. Paired gonads lie above or lateral to the midgut in most species. In mysids, the
ovaries are partially fused and linked by a cellular
bridge. Paired, or rarely single, reproductive ducts open
Structure of the antennal gland of a crayfish. [From Mantel and Farmer, 1983.]
784
A. P. Covich and J. H. Thorp
ventrally through simple gonopores (paired or single),
elevated papillae, or elaborate copulatory structures (as
in decapods). The location of the gonopore varies
greatly in entomostracans, whereas in malacostracans,
it opens on the sternite or coxae of the sixth thoracic
somite in females and at the eighth somite in males. Internal fertilization is the general rule, with males transferring sperm through a penis or specialized trunk appendages known as gonopods (in crayfish these are
highly modified pleopods).
Females of most species protect their embryos either by retaining them in an internal brood chamber
(e.g., in cladocerans) or external ovisac (e.g., in copepods) or by gluing them to certain appendages (decapods employ their abdominal pleopods). Mysids,
isopods, and amphipods safeguard their young until an
advanced stage in a ventral brooding shelf, the marsupium. Ovigerous female amphipods are readily recognizable when carrying eggs or young in late developmental stages because of their extended brood pouch
and its yellowish to grey coloration. Stream-dwelling
isopods apparently release their juveniles from the
brood chamber when at risk from fish predation, but
not when exposed to lower risk of predation by salamanders (Sparkes, 1996a,b).
The embryogeny of crustaceans includes modified
spiral, total cleavage, and gastrulation by invagination.
All taxa possess an initial nauplius stage in their development, but these larvae may be free swimming (e.g.,
in copepods) or enclosed in an egg (e.g., in decapods,
cladocerans, and peracaridans). Direct development
(i.e., without any external larval stages) characterizes
taxa such as cladocerans and peracaridans. The young
individuals appear morphologically identical to miniature adults. In contrast, indirect development with a
free nauplius followed by either distinctive stages of
metamorphosis or gradual development to an adult is
typical of most ostracodes, copepods, decapods (except
crayfish), and Branchiopoda (other than cladocerans).
Growth throughout the larval, juvenile, or adult
stages requires periodic shedding of the older, smaller
exoskeleton in a process termed molting or ecdysis.
Rapid expansion of the body occurs immediately after
the crustacean extracts itself from its old exoskeleton
and before the new shell hardens. The degree of expansion varies significantly among species, ranging from
8 – 9% in some mysids to 22% in many decapods, to as
high as 83% in certain cladocerans (Hartnoll, 1982).
Some taxa are characterized by indeterminate growth
(e.g., the cladoceran Daphnia and apparently most
crayfish species), while others reach a finite body size in
a fixed number of molts (e.g., in ostracodes, copepods,
and some isopods). The molt process is controlled by
hormones released by several organs. With a decline in
production of the molt-inhibiting hormones (normally
secreted by the cephalic X-organ and sinus gland complex), the Y-organs are no longer suppressed and can
begin secretion of ecdysone, a molt-stimulating hormone. The actual molt requires as little as a few minutes to as long as several hours in aquatic taxa. This period is extremely hazardous; the animal may die from
physiological stress or succumb when it is unable to extract a portion of its body from the old exoskeleton.
The newly molted individual is vulnerable to rapid colonization by epibionts and ectoparasites. Some species
of ciliated protozoans and sessile rotifers are well
adapted to quickly recolonize the clean surfaces of
newly molted isopods (Cook et al., 1998; Roberts and
Chubb, 1998). The temporarily soft shell of a recently
molted individual is extremely susceptible to predatory
attacks (and cannibalism). Some species may seek shelter prior to molting. The larger and older the individual
becomes, the longer it takes for the molting process to
be completed, and the longer is the exposure to possible
cannibalism and predation. Various environmental factors influence susceptibility of crustaceans to predators
after molting. For example, medium, and large-sized individuals experience lower risks of predation from salamanders when living in waters rich in calcium carbonate (tufa and travertine), because lime deposits on their
bodies apparently make them less vulnerable (Ruff and
Maier, 2000). Thus, although indeterminate growth
typically has limitations imposed by physiological
stress, parasite infections, epibiont loads, and predatory
pressures, each habitat may vary in how environmental
complexity influence rates of molting and survivorship.
C. Environmental Physiology
1. Salinity, Temperature, and pH Tolerances
A large number of crustacean species have adapted
to living completely in freshwater habitats (Table I) while
others spend only portions of their lives in these highly
variable habitats (e.g., Mantel and Farmer, 1983; Vernberg and Vernberg, 1983; Holsinger, 1994a; Wollheim
and Lovvorn, 1996). Branchiurans are distributed worldwide in both marine systems and freshwater habitats;
some species can tolerate a rapid shifting of distributions
from salt to freshwater and may even switch hosts from
marine to nonmarine fishes. Among the Peracarida, as
will be discussed later, some species are confined to
coastal inland waters and apparently are not completely
adapted to many of the relatively unbuffered, dilute waters found farther inland (Sutcliffe, 1971, 1974; Dormaar
and Corey, 1978). Amphipod species differ greatly in
their ability to osmoregulate in variable salinities. One estuarine species from the western coast of North America,
Corophium spinicorne, occurs typically in tidal fresh
19. Introduction to the Subphylum Crustacea
waters (Hutchinson, 1967). This species is reported to
have dispersed farther inland and now it occurs in fresh
waters at two sites along the Snake River, upstream of
Lewiston, Idaho (Lester and Clark, in press). This coastal
species apparently was transported upriver by barges and
may be widely distributed in the lower reaches of the
Snake and Columbia rivers. Some species, such as
Hyalella azteca, are found in saline lakes with salinities
up to 25 parts per thousand (Hammer et al., 1990).
Other amphipod species are sensitive to direct or indirect
effects of increased salinity (Wollheim and Lovvorn,
1996). Salt pollution from various sources can increase
hypo-osmotic stress and create adverse effects on amphipod metabolism (Koop and Grieshaber, 2000).
Temperature is an important regulator of metabolism and growth rates as well as survival. The upper
lethal temperature that limits survival varies greatly
among different species. Some crustacean species have
the highest known temperature tolerance among the
aquatic metazoans and can live in hot springs. For example, the ostracode Potamocypris, lives on algalcoated substrates in habitats that range from 30 to
54°C (Wickstrom and Castenholz, 1973), the pancarids
Thermosbaena mirabilis survives in waters of
45 – 48°C, and Thermobatynella adami lives in 55°C
water (Capart, 1951). Some isopods are found in
warm springs of the southwestern United States; for example, Thermosphaeroma thermophilum, an endangered species, and T. subequalum thrive in 34 – 35°C
pools (Bowman, 1981; Cole and Bane, 1978; Shuster,
1981, Manuel Molles, personal communication). Other
benthic crustaceans, such as the amphipod Hyallela
azteca, also live in hot springs (Richard Wiegert and
Dean Blinn, personal communication) with temperatures up to 33 – 34°C in Devils Hole, Nevada.
In general, individual growth rates of Hyalella
azteca and other crustaceans are affected directly and
indirectly by water temperatures. Variations in water
temperatures often result in seasonal movements by
differently aged individuals as they track changing temperatures at different depths. For example, field observations demonstrate that 20°C is a threshold for induction and termination of reproductive resting stages of
Hyalella azteca. This species moves from warmer, shallow littoral zones into colder, deeper water as individuals mature (Panov and McQueen, 1998). If water
depths decrease and water temperatures increase in response to both successional and climatic warming, it is
anticipated that benthic species such as Hyalella azteca
would likely have increased growth rates and earlier
breeding in shallow lakes, ponds and intermittent
streams. These shifts in water temperatures and habitat
fragmentation could affect intra- and interspecific competition and gene flow among benthic species, espe-
785
cially during prolonged droughts (Hogg and Williams,
1996; Covich et al., 1997; Hogg et al., 1998a). Other
amphipod species require colder, relatively constant
waters and are often associated with cold springs. For
example, Gammarus minus is rarely found in waters
warmer than 15°C and is absent in waters above 20°C
(Glazier et al., 1992). Monitoring these cool-water
habitats and their amphipod populations will provide
important information about the long-term effects of
climate on groundwater and surface-water temperatures (Wilhelm and Schindler 2000).
The widespread acidification of lakes and streams
has drawn attention to the importance of pH tolerances
of many organisms, especially the crustaceans. Because
calcification of the carapace immediately following molting is highly sensitive to low pH, changes in regional distributions among crustaceans may reflect regional levels
of acidification and natural differences in calcium concentrations (e.g., Nero and Schindler, 1983; Greenaway,
1985; Okland and Okland, 1985; Meyran, 1997, 1998;
Meyran et al., 1998). There are both lethal and sublethal
effects of increased acidity that alter ion regulation, especially by juveniles in soft-water lakes. Although many
studies have emphasized lentic species, there is evidence
that lotic species may have a narrower range of tolerance to low pH and that egg mortality may be important for some species. Crustaceans generally are good indicators of changes in pH values in well-defined
habitats. For example, a regional study documents that
Gammarus minus distributions in 32 springs in central
Pennsylvania are limited to waters with pH values of 6
and above, with most populations found in waters near
pH 7 and where temporal variation was minimal
(Glazier et al., 1992, Glazier and Sparks, 1997).
Although crayfish are generally thought to require
habitats with calcium concentrations in excess of
2 mg / L, low population densities of some species may
be maintained by extracting sufficient calcium from
their food. In adult Orconectes virilis, calcium uptake
is impaired in habitats below pH 5; survivorship is
greatly influenced by age and molt stage (Chapter 23).
2. Photoreception
Sensitivity to small differences in light intensities is
extremely well developed in crustaceans. Many groups
are able to locate microhabitats where they can optimize growth rates and reproduction (Hutchinson, 1967,
1993). Crustaceans have evolved receptors to detect
even slight differences in light intensities, especially in
deep, thermally stratified waters. Marine and freshwater zooplankton exploit thermal gradients by migrating
daily and seasonally in both vertical and horizontal patterns to obtain their preferred conditions. The adaptive
value of this migration for zooplankton is thought to
786
A. P. Covich and J. H. Thorp
be a combination of: (1) optimizing growth through
reducing metabolic costs by remaining in cool waters;
(2) maximizing growth through grazing at times when
algae have the highest food value; and (3) avoiding exposure to visual predators. The relative importance of
these variables can change seasonally and shift the exact
timing of migration to maximize net energy gain under
different conditions. When algal food resources are
scarce, zooplankton may ascend “early” from deep cool
waters and begin grazing 1 – 2 h before sunset, thereby
gaining access to limited food supplies before competitors, but at a higher metabolic cost and risk of predation (Enright, 1977, 1979; Gliwicz, 1986). In arctic
lakes and ponds, crustacean zooplankton are exposed
to continuous daylight during most of their growing
season and do not vertically migrate on a daily basis.
However, they maintain phototactic reactions at approximately the same threshold values; their photosensitivity remains similar to temperate species (Buchanan
and Haney, 1980). Controversy exists regarding how to
evaluate the adaptive significance of variable migratory
responses to different light intensities in the presence of
predators (see chapters 21 and 22).
Many decapods are nocturnal and avoid sight-feeding predators by remaining inactive and undercover
during the day. They have sensitive photoreception and
synchonize their activity cycles within their population
and thus increase their nighttime densities and reduce
their individual vulnerability to predators. Many species
of benthic amphipods and isopods seek dark microhabitats such as beneath leaf litter during daylight to minimize exposure to visual predators. This normal avoidance behavior of bright habitats by amphipods and
isopods, however, can be affected by endoparasites. Infected crustacean hosts increase their daytime activity
and can thereby increase their vulnerability to predators. This predation often continues the life cycle for the
parasite with the predatory fish or bird serving as a second intermediate host (e.g., Helluy and Holmes, 1990;
Maynard et al., 1998). How photoreception and activity cycles of some crustacean species are altered by specific parasites is an active area of research.
3. Chemoreception
Crustaceans also have very well-developed chemosensory systems that allow individuals to locate food and
mates while avoiding predators (Ache, 1982; Atema,
1988; Zimmer-Faust, 1989). Crustaceans demonstrate
highly specific responses to pheromones (Dahl et al,.
1970; Dunham, 1978; Dahl et al., 1998; Dahl, 1998;
Wooster, 1998, see chapters 20 – 23). Hydrodynamics of
stream flow and lake currents clearly mediate the
effectiveness of diffuse chemicals, but the persistent unidirectional nature of many signals does provide a basis
for orientation and communication. Laboratory and
field studies demonstrate that chemical communication
between crustacean predators such as crayfish and their
gastropod prey can influence prey life histories (Crowl
and Covich, 1990). Crustacean prey, such as Gammarus minus, respond differently to chemical cues released from injured conspecific individuals relative to
chemical cues released from other species such as sympatric
isopods,
Lirceus
fontinalis
(Wisenden
et al.,1999). Gammarus lacustris responds to alarm signals from fish predators (northern pike, Esox lucius),
but not to dragonfly larvae (Aeshna eremita), apparently because fish are more effective predators (Wudkevich et al., 1997). Similarly, in laboratory studies
Lirceus fontinalis reduces their activity in the presence
of alarm substances from fish mucus of predatory green
sunfish (Lepomis cyanellus) but not from grazing fish,
stonerollers (Campostoma anomalum) (Short and
Holomuzki, 1992). Within the boundary layer of
stream flow, chemoreception can be highly effective as a
means for two-dimensional orientation. For organisms
drifting downstream or migrating upstream, chemical
cues may be extremely important. The relative size of
the organism to the depth of the boundary layer is an
important ratio (Dodds, 1990) that influences effectiveness of chemical communication.
During periods of stable-flow regimes in streams,
crustaceans may depend on chemical cues as well as
other sources of information to regulate daily and seasonal patterns of movement, to select foods or mates,
and to avoid predators. As pointed out in chapters 21
and 23, there are recent studies on the effectiveness of
chemical communication in crustaceans, and the situation is similar for other groups (Dodson et al., 1994).
4. Mechanoreception
The ability to orient into currents (positive rheotaxis) is a common trait in many freshwater crustaceans.
These physical, hydrodynamic cues are often associated
with chemical cues that stimulate directed movement.
Crustaceans have many types of setae, which function
as mechanical and chemical sensors (Bush and Laverack, 1982). These external receptors are typically positioned near the antennae and the mandibles. However,
there is a wide range of other locations so that information can be obtained from several directions simultaneously. The functional anatomy of these mechano-receptors is well studied in decapods. Behaviors associated
with frequent cleaning activities are not as thoroughly
studied in freshwater crustaceans as in marine taxa nor
are there many studies on behavioral responses to
changes in current speed (see Chapter 23 for further discussion). Behavioral studies of copepods have emphasized the importance of mechano-receptors for avoidance of predators (see review in Chapter 22), but
relatively little is known for other groups.
19. Introduction to the Subphylum Crustacea
5. Thermoreception
In warm waters rates of metabolism increase and
the need for dissolved oxygen also increases. However,
concentrations of dissolved oxygen decrease in warm
waters and low-oxygen stress can cause mortality. This
low-oxygen stress can be avoided if individuals move
to certain microhabitats (e.g., groundwater seeps and
deeper, stratified waters in summer). Thus, sensing differences in relative temperatures is an important ability
for crustaceans to locate optimal microhabitats for
growth and reproduction. Many species can orient in
thermal gradients and combine information on temperature with simultaneous inputs from photo- and
chemoreceptors (Ache, 1982). Freshwater crustaceans
have distinct thermal preferences (e.g., Cheper, 1980;
Oberlin and Blinn, 1997) and considerable information
is available for crayfish (as discussed in Chapter 23). As
mentioned previously, the interactions between thermal
cues and light intensity are often associated with vertical migrations of zooplankton. In these behavioral responses to light and temperature gradients, there is
growing evidence that zooplankton may also use chemical cues to orient themselves and to avoid predators
(see chapters 21 and 22).
III. ECOLOGY AND EVOLUTION
OF SELECTED CRUSTACEA
A. General Relationships
Evolution within the Crustacea has resulted in a
very large number of different species. The fossil record
demonstrates that both freshwater and marine crustaceans existed in the early Paleozoic and three main
lines evolved from the monophyletic origin of arthropods (Schram, 1982; Emerson and Schram, 1990). Almost all of the crustaceans, that have adapted to freshwater environments, are thought to have evolved
directly from marine ancestors without having gone
through a phase of terrestrial adaptation (Hutchinson,
1967; Notenboom, 1991; Hutchinson, 1993; Holsinger,
1994a; Lee and Bell, 1999; Strayer, 1999).
The class Remipedia is a group of blind crustaceans that occur only in unique coastal (anchialine)
cave environments (Yager, 1981, 1987; Schram, 1986;
Schram et al., 1986; Felgenhauer et al., 1992; Carpenter, 1999). These crustaceans have a long trunk with
32 unfused segments and each segment bears a pair of
similar biramous appendages. Remipedes can tolerate
saline waters with very little dissolved oxygen and
they apparently feed in overlying, well-oxygenated
freshwater lens. This unique ecological salinity range
gives them some advantages over other crustaceans
living in marine polyhaline coastal cave environments.
787
Remipedes are considered to be primitive crustaceans,
and their evolution and biogeography have undergone
active study since the relatively recent discovery of
living representatives, primarily in the Caribbean region (Bahamas, West Indies, and in karst wells or
“cenotes” of Yucatan, Mexico) but also from the
Canary Islands. There are known to be fewer than a
dozen species in two families. Their structural
anatomy is better known (from transmission and
scanning electron microscopic studies) than their
functional anatomy, because they have not been successfully reared in the laboratory and studied experimentally (Felgenhauer et al., 1992). Other freshwater
crustaceans also occur in anchialine caves and other
complex hypogean habitats as discussed below (Stock
and Iliffe, 1990; Holsinger, 1993, 1994a; Holsinger
et al., 1994; Notenboom et al., 1994; Sket, 1996;
Danielopol et al., 2000). Macrocrustaceans adapted
to caves, artesian wells, and other groundwater habitats include a large number of amphipods, isopods,
mysids, and decapods (Chapter 23) that inhabit these
isolated inland waters (Holsinger, 1972; Holsinger
and Longley, 1980; Abele, 1982; Bousfield, 1983;
Barr and Holsinger, 1985; Holsinger, 1986, 1988,
1989, 1991, 1994a, b; Stock, 1995).
Although most benthic crustaceans are small and
difficult to observe without intensive sampling, there
are very large-sized crustaceans living in small and
large rivers. Macrobrachium are the largest (20 cm
carapace length) freshwater crustaceans in North
America and are distributed primarily in coastal rivers
along the Gulf of Mexico (Chapter 23). They usually
have amphidromous life cycles that limit their populations to coastal areas connected to brackish and marine
waters where they spend part of their series of complex
larval stages. Although more than 90 species occur in
the tropics, there are only six species of these palaemonid shrimp in North America. However, their riverine migrations and long-term survival are vulnerable
because of barriers created by impoundments, overharvesting, and water pollution. Only M. ohione is
known from more interior rivers and its populations
are thought to be declining (Bowles et al., 2000).
1. Branchiura
The subclass Branchiura is widely distributed
throughout the world and is comprised of about 150
species and four genera that are ectoparasites on fish.
Although there are many freshwater species (Yamaguti,
1963; Cressey, 1972; Margolis and Kabata, 1988),
only the genus Argulus is typically found in North
American freshwater habitats (McLaughlin, 1980;
Abele, 1982; Fitzpatrick, 1983; Schram, 1986). Some
23 North American species are encountered; three are
found in western and 19 in eastern North America.
788
A. P. Covich and J. H. Thorp
The largest number of species occurs along the Gulf of
Mexico. Only a few are very restricted in their geographic ranges, while others are widely distributed; for
example, A. maculosus occurs from New York to
Michigan and down the length of the Mississippi River
to Louisiana.
For many years, the Branchiura, or “fish lice,”
were classified as a group of Copepoda (Wilson, 1944)
and earlier as Branchiopoda, but their separation as a
distinct taxon is now widely accepted. Unlike copepods, the argulids have a pair of compound eyes. The
Branchiura also differ from parasitic copepods in having several other unique structures that will be described. The branchiurans have undergone major modifications of the mandibles and associated feeding
appendages. The mandibles typically form a pair of
transversely toothed hooks between the labium and
labrum, and in Argulus, the mandibles are incorporated into the specialized proboscis (McLaughlin,
1982). The maxillules are large suckers that are characteristic of adult branchiurans (Fig. 6). In Argulus, a
sheathed, hollow spine is used to pierce the skin of the
host. Some species are blood suckers while others feed
on extracellular fluids or mucous. Once engorged, they
can wait 2 – 3 weeks between meals.
Some reproductive and larval behaviors are unique
to Branchiura. In contrast to copepods, female
Branchiura do not carry eggs in external ovisacs but,
instead, fasten them in rows to rocks and other objects.
The actively swimming larvae attach themselves within
FIGURE 6
External anatomy of an ectoparasitic “fish lice”
(Branchiura: Argulus).
the gill chambers, mouth, or on the outer surfaces of
fishes using small, specialized antennal hooks and maxillules. After 4 – 5 weeks of development as ectoparasites, they become adults that again may actively swim
(actually more like somersaulting through the water) to
find mates and other hosts. Adult females are highly
motile in seeking egg-laying sites. They apparently are
opportunistic in selecting host species of fish (Lamarre
and Cochran, 1992; Shafir and Oldewage, 1992). The
mucous on fish surfaces does not interfere with attachment and low-level, short-term infection may not directly affect the fish host (Nolan et al., 2000).
2. Peracarida
The superorder Peracarida has a relatively low percentage of taxa living in inland waters. The three orders that comprise the Peracarida are the Amphipoda,
Isopoda, and Mysidacea. These orders are commonly
known as “scuds” (also “sideswimmers”), “sow bugs,”
and “opossum shrimp,” respectively (Figs. 7 – 9). Members of a fourth order, the Thermosbaenacea, with
some species living in thermal springs and others in
fresh or saline groundwaters, have strong peracarid
affinities but are generally considered to belong to the
superorder Pancarida. Only one taxon, Monodella texana, lives in North American inland waters (Maguire,
1965; Stock and Longley, 1981). Most carcinologists
exclude the Thermosbaenacea from the Peracarida because they brood their eggs dorsally under the carapace
rather than in a brood pouch, a unique feature of the
peracarids (Schram, 1982 1986). Within the Mysidacea, most of the 780 species are marine; worldwide,
only 25 species occur in freshwater, and 18 additional
species live in freshwater caves (Abele, 1982; Schmitz,
1992).
The Amphipoda is the largest peracarid order with
some 6000 known species in more than 100 families
(Schmitz, 1992). Amphipods are mostly small (5 –
15 mm in length), free-living benthic organisms
although some are pelagic and a few are semiterrestrial.
There are three large families, Gammaridae, Crangonyctidae and Hyalellidae and worldwide, there are
approximately 800 gammarid amphipods (Abele,
1982; Pennak, 1989). In North America, these amphipods are represented mainly by six families (Table
II) and about 175 species that occupy surface waters.
Another 10 families of amphipods (composed of 28
genera and approximately 125 – 160 species) occupy
subterranean waters (Holsinger, 1986, 1994a).
Compared to the Amphipoda, the Isopoda is a
much less species-rich order but with endemic species
often restricted to specific cave systems and other subterranean habitats. The Asellidae is the largest isopod
family in North America with about 115 aquatic
19. Introduction to the Subphylum Crustacea
789
FIGURE 7 External side view of generalized freshwater gammarid amphipod: 1, head; 2, antenna 1; 3,
antenna 2; 4, mouth parts; 5, pereonites 1 – 7; 6, pereopods 1 – 7; 7, pleonites 1 – 3; 8, pleopods 1 – 3; 9,
uronites 1 – 3; 10, uropods 1 – 3; and 11, telson (redrawn from Holsinger, 1972). Top view of telsons: 12,
Gammarus; and 13, Crangonyx.
species, of which approximately 65 are cave-dwelling
species. The other families only contain 10 species in
surface and six in subsurface waters of North America
(Sket, 1999a). The relative scarcity of isopods in many
freshwater habitats and their wider occurrence in hypogean waters and thermal springs may be related to
their lack of competitive ability and more restricted
timing of reproduction relative to amphipods and other
benthic invertebrates (Bowman, 1981; Graça et al.,
FIGURE 8 Diagram of top view of isopod Caecidotea (drawn
without setation).
1994a, b). In Europe, Asellus aquaticus generally
occurs in the lower reachers of river drainages while
Gammarus pulex is more common in the upper
reaches, but their competitive interactions and susceptibility to predation are not completely documented.
Isopods, especially juveniles, are sensitive to low oxygen concentrations, toxic metals, and organic enrichment of habitats. Ratios of widespread species of
isopods and amphipods (such as Asellus and Gammarus) are used to compare regional water-quality
conditions (Naylor et al., 1990; Maltby, 1991, 1995;
Whitehurst, 1991; Mullis et al., 1996). Examples of
widely distributed genera of isopods, such as Caecidotea Lirceus, and Asellus, are well studied and often
compared with associated amphipod species in surface
and sub-surface waters.
The Peracarida share several evolutionary similarities that demonstrate a common lineage. For example,
all have modified the ancestral first thoracic leg into a
mouth part (the maxilliped) and retain seven pairs of
thoracic legs for movement. The first two pairs of these
thoracic legs (the gnathopods) are specialized for grasping food while the remaining five pairs of legs (the
pereiopods) lack any specialized grasping (chelate) appendages. As mentioned previously, peracarid females
carry their eggs and young until a relatively advanced
stage of development in a specialized ventral brooding
chamber, the marsupium. Another character used for
separating the peracarids from most other crustaceans
is the highly developed lacinia mobilis, a small toothed
process that articulates with the incisor process. A
row of spines commonly separates the lacinia mobilis
and the molar process (McLaughlin, 1982). Several
phylogenetic relationships have been proposed for the
790
A. P. Covich and J. H. Thorp
FIGURE 9
Diagram of side view of opossum shrimp, Mysis.
evolution of the Peracarida (see Schram, 1982, 1986,
for review) based on how the fossil record is interpreted relative to modern morphological criteria. Many
workers view the Isopoda and Amphipoda as derived
from a common mysidlike stock. Watling (1981) proposed a different arrangement that is more consistent
with the known, but incomplete, fossil record (Fig. 10).
Most workers agree that the amphipods and mysids are
more closely related to each other than to the isopods
(Schram, 1982, 1986; Sieg, 1983).
Evolutionary relationships among species and subpopulations of widely distributed species are active areas of research (as discussed below) and new molecular
techniques are providing insights regarding differences
in rates of speciation. For example, studies using molecular time scales based on allozyme analysis suggest
that the divergence of several amphipod lineages that
gave rise to the three main species of Pontoporeia
(P. affinis, P. hoyi, and P. femorata) is likely to have
occurred over tens of millions of years. The divergence
of Mysis relicta from its likely marine congeners apparently dates from the Tertiary (Vainola and Varvio,
1989; Vainola, 1990).
TABLE II Representative Taxa Found in North
American Fresh Waters
Superorder Peracarida
Order Amphipoda
Family Crangonyctidae
Crangonyx, Stygobromus
Family Gammaridae
Gammarus minus, G. lacustris
Family Hyalellidae
Hyalella azteca, H. montezuma
Family Pontoporeiidae
Monoporeia affinis
Order Isopoda
Family Asellidae
Asellus, Caecidotea, Lirceus
Order Mysidacea
Family Mysidae
Mysis relicta, M. littoralis, Neomysis mercedis,
Taphromysis louisiannae
FIGURE 10
Alternative phylogenetic interpretations of the Peracarida: (A) widely accepted relationship of Mysidacea as the main
stem from which two divergent lines evolved, one to Isopoda and the
other to Amphipoda; (B) another view derived from Watling (1981)
with independent lines of development for each major group. The orders Cumacea, Tanaidacea, and Spelaeogripacea are marine peracarids, while the order Thermosbaenacea is often considered a pancarid group closely related to the peracarids. [From Schram, 1982.]
19. Introduction to the Subphylum Crustacea
B. Ecological Distributions and Interactions
1. Habitats
The widespread distribution of Branchiura suggests
a lack of any specific pH or salinity requirements, but
detailed studies of these ecological relationships are
lacking. Temperature influences the rate of hatching for
eggs of different species with a range of 12 – 30 days
being required (Yamaguti, 1963). During the freeswimming stage Argulus may aggregate in a narrow
zone of temperature and light conditions where encounters with host fishes can be increased. Argulus has
no strict host specificity and is capable of attaching to
both freshwater and marine telosts.
Most peracarid taxa that have fully adapted to the
physiological and ecological conditions of inland
waters share five ecological similarities in that they: (1)
are typically restricted to permanent bodies of water
that are relatively cool, clean, and well oxygenated; (2)
have distinct behavioral patterns of vertical migration
(e.g., among mysids and two species of amphipods;
these are similar to daily migration in other groups of
crustacean zooplankton, as discussed previously and in
chapters 21 and 22); (3) have relatively limited ability
to move upstream (positive rheotaxis is well studied in
amphipods and isopods) or drift downstream in the
current under specific ecological conditions (as do
many of the lotic insects discussed in Chapter 18); (4)
obtain much of their energy while feeding on the bottom substrates (even if they also feed while in the nekton on suspended algae and smaller taxa of zooplankton); and (5) serve as important prey to a large number
of predatory fishes (also discussed in chapters 21 – 23).
Some taxa also have various ways of inflicting damage
to predatory populations either by being direct parasites on predators or by competing with young stages
of those predators for the same prey (such as Mysis
feeding on plankton) that would otherwise be potentially available to fish consumers.
In contrast to other crustaceans, most of these peracarid species typically lack a diapause phase and are
missing any distinct adaptation for avoiding desiccation. Thus, their chances for passive dispersal (e.g., being carried from one habitat to another by ducks or
other migratory animals or being carried by the wind
during storms) are low in comparison with cladocerans, copepods, or ostracodes (Hairston and Caceres,
1996; Hairston and Bohonak, 1998; Bohonak, 1999).
The peracarids also lack any strong active dispersal
ability. They generally do not move upstream against
strong current (Olyslager and Williams, 1993) so
their migration throughout river-drainage networks is
restricted, and populations apparently remain more or
less in the same locality for long periods. Nonetheless,
791
as discussed later, some taxa are very widely distributed
and their patterns of distribution suggest a long history
of slow dispersal and persistence.
There are a few very widely distributed amphipod
species in surface waters. Some surface-dwelling and
subsurface species have been well studied ecologically
so that comparisons of population densities and life
histories provide important information for their conservation and as indicators of water quality (e.g.,
Mullis et al., 1996; Mosslacher, 1998; Knapp and
Fong, 1999). The most widely distributed and diverse
genera of cave-inhabiting amphipods are Stygobromus
with approximiately 180 species and Crangonyx with
14 species (Peck, 1998). No doubt, many species remain undescribed in caves, hyporeheic zones, deep
lakes, and even in well-studied, shallow-water habitats.
The continued discovery of new species, and the recent
applications of new techniques for genetic analysis of
previously described taxa, suggest that many more genetically distinct populations occur than was first expected (Hogg et al., 1998b). Many species of amphipods and isopods are exceptionally well adapted to
live in surface waters as well as groundwater habitats
and caves (Holsinger, 1972, 1986, 1988; Stanford and
Ward, 1988; Fong, 1989; Culver et al., 2000). In cave
ecosystems, a single species of amphipod may dominate
a relatively simple food web based on fine organic particulates (e.g., Drost and Blinn, 1997). The microbiogeographical distributions and evolution of relatively
common species such as Gammarus minus are under
intensive study (Kane et al., 1992; Sarbu et al., 1993;
Culver et al., 1994; Culver and Fong, 1994; Fong and
Culver, 1994; Culver et al., 1995; Kane et al., 1995).
As discussed below, other intensive studies have focused on the evolution of subpopulations of Hyalella
azteca; some apparently have formed distinct species in
certain surface-water habitats (Thomas et al., 1997;
McPeek and Wellborn, 1998).
Ecologically, most of the amphipods and isopods
are photonegative, positively rheotactic, thigmotactic,
cold stenotherms (i.e., restricted to relatively constant,
cold waters where they avoid bright light by moving
into the current and into crevices or under stones,
leaves and roots). In complex substrates, they are less
exposed to predators such as fish and crayfish. Typically, where refugia from predation exist, the peracarids occur at high densities in small, permanent,
spring-fed streams, seeps, ponds, or sloughs (e.g., Allee,
1914, 1929; Juday and Birge, 1927). In isolated springs
their populations sometimes genetically diverge from
those in other springs (Gooch and Glazier, 1991;
Glazier et al., 1992; Culver et al., 1994; Glazier, 1999).
Some species of Gammarus reach densities of thousands of individuals per square meter where detrital
792
A. P. Covich and J. H. Thorp
food and cover are abundant and predators are few
or absent.
A relatively smaller number of isopod species have
apparently adapted to live in intermittent or permanent
surface waters but distributional data are scarce
(Richardson, 1905; Hubricht and Mackin, 1949;
Williams, 1970; Ellis, 1971). Certain species are common in subsurface habitats. For example, the subterranean isopod Caecidotea tridentata is relatively abundant in groundwaters of the Konza prairie (Edler and
Dodds, 1996). The most diverse genus of cave-inhabiting aquatic isopods is Caecidotea with more than 56
species (Peck, 1998). For most genera of isopods the
number of described species found in groundwater
habitats and caves of North America ranges from one
to five (Holsinger, 1988; Fong and Culver, 1994; Culver et al., 1995; Peck, 1998). Only a few known species
tolerate the low oxygen concentrations found in polluted rivers and groundwaters (Williams, 1970; Strayer
et al., 1997; Henry and Danielopol, 1998; Mosslacher,
1998; Hervant et al., 1999; Malard and Hervant,
1999) or the stress of high temperatures (Dadswell,
1974). For example, some isopods (Caecidotea recurvata) survived in cave pools only slightly polluted by
septic tank effluents, but were absent from highly
polluted pools (Simon and Buikema, 1997). There is
concern that any level on increased food input by organic pollution could result in competitive displacement of subsurface species by surface-dwelling species
that are better adapted to use higher concentrations of
food (Sket, 1999b).
The order Mysidacea has only a single family and
one very important species, Mysis relicta (Fig. 9),
which occurs in cold, deep lakes as well as in shallow
brackish ponds along the arctic coasts. A second
species, Neomysis mercedis, has a more limited, coastal
distribution but plays a significant role in some large
lakes such as Lake Washington (Murtaugh, 1989). Mysis relicta, the “opossum shrimp,” is not a true shrimp
(or decapod as defined in Chapter 23), but is an active
swimmer and can make long daily trips up and down
the water column of deep, well-oxygenated lakes
(Beeton and Bowers, 1982; Shea and Makarewicz,
1989). A third species, Taphromysis louisianae, lives in
roadside pools in Louisiana and Texas (Banner, 1953;
Pennak, 1989).
Mysids have a holarctic distribution; they occur in
oligotrophic lakes of northern regions of North America and in northern Europe, but are known to tolerate
relatively high temperatures and periods of low oxygen
at least for short periods (Dadswell, 1974; Sherman
et al., 1987). As will be discussed later, their introduction into many western lakes has created several problems for fisheries managers (Martinez and Bergersen,
1989). Their natural limits of distribution apparently
have been influenced by opportunities for migration
during the Pleistocene and possibly by biotic interactions (Dadswell, 1974).
2. Food Resources
All three peracaridian orders are similar in the
range of foraging behaviors used in obtaining food resources. Juveniles are typically dependent on microbial
foods such as algae and bacteria associated with either
periphyton or aquatic plants in brightly illuminated
habitats. They also consume dead organic matter in
forested streams, ponds, and lakes where bacteria and
fungi living on the detritus provide essential protein for
amphipods and isopods (Barlocher and Kendrick,
1973, 1975; Smock and Stoneburner, 1980; Smock and
Harlowe, 1983; Findlay et al., 1984, 1986; MacNeil
et al., 1997). Adults are not limited to grazing or detritivory because they broaden their food niche to include
larger food items and are predatory (Schwartz, 1992;
MacNeil et al., 1999b). Some pelagic or nektoplanktonic amphipods in fishless lakes, ponds, and wetlands
consume phytoplankton and zooplankton (Anderson
and Raasveldt, 1974; Dehdashti and Blinn, 1991).
Adults of well-studied species, such as Gammarus minus, are known to depend primarily on detritus when
in surface waters but cave populations of this species
are more flexible in their diet (Culver et al., 1995). In
addition to the many plant, animal, and detrital foods
used by different species of Gammarus, intraguild predation and cannibalism are also thought to be relatively common in surface waters (Dick, 1995; MacNeil
et al., 1997, 1999b). Amphipods become opportunistic
scavengers, predators, and omnivores, depending on
food quality and which foods are most available (Delong et al., 1993). Given their wide range of foods, it is
clear that different modes of feeding are used to exploit
these different resources (e.g., Crouau, 1989). These
generalized consumers can reach high population
densities by deriving their energy from many sources.
Resource competition among guild members can be
intense if access to suspended foods is space-limited, as
may occur in soft-bottom streams among filter-feeding
species that require hard surfaces for attachment or
perching. Interference competition and aggressive
behavior can also occur among species that differ in aggressiveness and mobility (e.g., Dick et al., 1995;
Haden et al., 1999).
Laboratory and field studies of production have focused on juvenile and adult growth rates that result
from feeding on different types and qualities of foods
at widely different locations (Waters and Hokenstrom,
1980; Marchant and Hynes, 1981a; Sutcliffe et al.,
1981; Delong et al., 1993; France, 1992, 1993). Only a
19. Introduction to the Subphylum Crustacea
few studies have considered the role of secondary plant
chemicals as “defensive” compounds for aquatic plants
to minimize effects of grazers. Concentrations of tannins, and other inhibitory chemicals are known to differ among various macrophytes, and some compounds
influence crustacean grazing preferences (e.g., Newman
et al., 1990, 1992). Plant foods that contain a high percentage of cellulose are often considered to be of relatively low quality for grazers and detritivores (Newman, 1991). However, some amphipods (Monk, 1977)
and mysids (Friesen et al., 1986) have digestive enzymes that hydrolyze cellulose in vitro. These cellulases, however, are apparently confined to degrading
small particles in the gut rather than whole plant cell
walls that are ingested during grazing. Microbial
breakdown of ingested cellulose within the gut is generally the mechanism used for digesting this refractory
material and requires development of a complex community of various microorganisms within the gut.
Given that crustaceans molt periodically and would
need to re-establish this microbial community in their
new gut lining, the evolution of cellulases would be an
important adaptation for expanding use of detrital
food resources.
3. Vertical Migrations and Feeding
Amphipods and isopods are often confined to burrowing and feeding on the bottoms of streams and
lakes, although species like Monoporeia affinis (Pontoporeia affinis) move from their benthic foraging areas
during the day to nektonic, open-water foraging at
night (Marzolf, 1965; Winnell and White, 1984; Donner et al., 1987). These actively swimming amphipods
consume a wide range of particles while on the sediment surface, although the exact nature of the food is
not completely known for those species found in the
Laurentian Great Lakes or other large basins (Dermott
and Corning, 1988; Lopez and Elmgren, 1989; Nalepa
et al., 2000). The seasonal deposition of diatoms and
other phytoplankton from the surface waters provides
important food resources for bottom-dwelling amphipods (Johnson and Wiederholm, 1989; Hill et al.,
1992; Johnson and Wiederholm, 1992; Sly and
Christie, 1992; Goedkoop and Johnson, 1994).
The vertical movements of amphipods, such as
Monoporeia affinis, in deep lakes overlap those of
mysids with whom they may share food resources.
Mysids provide an intriguing example of vertical migration and complex foraging dynamics (Chess and
Stanford, 1998, 1999; Rudstam et al., 1999; Spencer et
al., 1999). Mysis relicta regulates nocturnal feeding
patterns by responding to changing light conditions
(Johannsson et al., 1994). These very fast, active swimmers have well-developed eyes for precise photorecep-
793
tion. They rely on daily changes in intensity and
quality of light (that penetrates through the photic
zone) to determine when to begin swimming up
through the water column from the dimly lit, deeper
waters and to begin feeding on small zooplankton and
phytoplankton (Morgan et al., 1978; Cooper and
Goldman, 1982; Folt et al., 1982).
By precisely regulating their travel times, migrating
crustaceans reduce their exposure to fish predators and
still obtain the food they need to grow and reproduce.
Both Mysis and Monoporeia have potential fish predators that feed on benthic and open-water habitats so
that minimizing exposures to a wide range of predators,
while also sometimes competing with them for zooplankton prey, is a complex process (McDonald et al.,
1990; Johannsson, 1992; Johannsson et al., 1994; Johannsson, 1995; Wilhelm, 1996). As mentioned previously, those zooplankton that seek deeper, cooler waters
during the day not only avoid brightly lit waters and exposure to visual predators, but they also lower metabolic rates while digesting the food that they obtained
the previous night (Chipps, 1998; Rudstam et al.,
1999). Depending on the temperatures of the upper
strata, Mysis migrates faster and closer to the surface
than does Monoporeia (Wells, 1960). Because Mysis has
less tolerance for low concentrations of dissolved oxygen in the deeper strata than does Monoporeia, it may
not spend much time in those deep layers during the
later portion of the growing season. Decomposition of
organic matter in the deep hypolimnion depletes dissolved oxygen if the upper waters are highly productive
and dead matter falls into these deep waters. Of considerable importance is the observation that Mysis can eat
Monoporeia (Parker, 1980). The spatial refuge from
mysid predation for amphipods then is dependent on
the length of time that a lake remains thermally stratified, its level of productivity, and its bottom water temperature, because these factors determine both the
concentration of dissolved oxygen in the deepest strata
and the presence of various benthic predators.
Because mysids are consumed by a number of
game fishes, there was a period of widespread, intentional introduction of Mysis relicta into many deep
lakes in the western and northern United States (Martinez and Bergersen, 1989; Spencer et al., 1999) and in
western Canada (Lasenby et al., 1986). As often happens with introductions of species into habitats where
they do not naturally occur, the foodweb dynamics did
not develop as expected. Of the 134 recorded introductions into the lakes of western North America, only 45
became directly established and eight others became indirectly established through passive migrations to other
lakes (Martinez and Bergersen, 1989). In some lakes,
instead of providing more food for predatory fishes
794
A. P. Covich and J. H. Thorp
such as lake trout and coregonids, the mysids ate many
of the same zooplankton species that the juvenile fish
had previously consumed (Lasenby et al., 1986). Thus,
the mysids began competing with fish for zooplankton
prey (Cooper and Goldman, 1982; Spencer et al.,
1991, 1999). In these lakes, the mysids did not migrate
high enough into the surface waters where the visually
oriented fish could capture them. In the spring and
summer, mysids avoided the warmer, upper, dimly lit
surface waters and thus escaped predation; furthermore, they intercepted the upwardly moving zooplankton at midwater depths before the zooplankton could
be consumed by fish. Even more unexpected was the
predation by Mysis relicta on newly hatched larval fish
(Seale and Binowski, 1988).
An unusual example of finding food occurs with
the amphipod Hyalella montezuma. This species migrates to the surface waters of a large karst sink hole
(Montezuma Well, Arizona) and filters out organic material entrapped on the surface film (Cole and Watkins,
1977; Blinn and Johnson, 1982; Blinn et al., 1987;
Wagner and Blinn, 1987). Its twilight migration is followed by a predatory leech, which swims up to feed on
the dense concentration of amphipods and other prey
(Blinn et al., 1988, 1990; Blinn and Davies, 1990;
McLoughlin et al., 1999); no fish predators occur in
this sink-hole lake. In contrast, Hyalella azteca, which
coexists with H. montezuma in Montezuma Well, does
not filter feed or migrate to the surface. In comparative
studies of Devils Hole, Nevada, Dean Blinn and Kevin
Wilson (personal communication) observed that H.
azteca migrate horizontally from the shoreline to the
open-water column at night, which is the reverse of
what occurs in Montezuma Well (Blinn et al.,1987). In
fishless lakes, ponds, and wetlands, other amphipods,
such as Gammarus lacutris and G. pulex, are known to
feed in the water column as well as on the bottom
(Anderson and Raasveldt, 1974).
Other well-studied examples of complex vertical
migrations of predatory macro-crustacean feeding on
zooplankton occur among species living in very deep
lakes and are similar to massive aggregations of Mysis
relicta in North American lakes (Lasenby et al., 1986).
As discussed below, Macrohectopus branickii in ancient Lake Baikal has dynamics and functions similar
to North American mysids (Rudstamet al., 1992;
Melnik et al., 1993).
4. Responses to Water Quality
The general requirement for habitats with relatively high concentrations of dissolved oxygen by most
peracarid crustaceans usually limits them to clean,
cold waters especially, during their early life histories.
Some species of amphipods are sensitive to changes in
temperature, stream discharge, and predators in their
choice of microhabitats through their onset of downstream drift responses and ability to move upstream in
search of appropriate conditions (Waters and Hokenstrom, 1980; Williams and Moore, 1985). In productive habitats crustaceans can deposit excess energy in
the form of lipid which they store internally (Hill et al.,
1992; Adare and Lasenby, 1994; Cavaletto et al., 1996;
Nalepa et al., 2000). Because they feed on a variety of
materials and incorporate chemical constituents from
their food into their bodies (often in their lipids), these
organisms are important in research on water-quality
monitoring (Conlan, 1994; Landrum and Nalepa,
1998; Song and Breslin, 1998, 1999). Many species
also move over relatively large areas and, thus, are exposed to a wide array of microhabitats that may contain specific toxins. The ease of collecting amphipods
and isopods from a broad range of habitats and their
hardiness in laboratory mesocosms make these crustaceans ideal candidates for monitoring pollution. They
are especially sensitive to copper (even from copper
pipes in laboratory plumbing) and a number of other
toxic heavy metals (Thybaud and LeBrac, 1988;
Borgmann and Munawar, 1989). They can be used as
“sentinels” of chronic exposure to low concentrations
of toxins or as indicators of acute episodes of toxic
spills that might otherwise go unmeasured by routine
sampling of streams and lakes.
Accidental spills of toxins can have major impacts
on survival of crustaceans and their ecological roles in
maintaining high water quality. Experimental studies
demonstrate the importance of detrital processing by
benthic crustaceans through comparisons of reference
“control” streams with “treated” or disturbed streams
(e.g., Newman et al., 1987). Mysids also have become
important organisms for monitoring the movement of
toxins through foodwebs in large lakes (Evans et al.,
1982). The presence or absence of Mysis relicta in
different lakes is important for understanding how
toxins such as mercury accumulate in top fish predators because of the major role these crustaceans play in
fish production (Cabana et al., 1994; Branstrator et al.,
2000).
C. Life-History Traits
1. Longevity
Although most small species appear to be “annuals” and may complete their life cycle within a single
year, larger amphipod species such as Monoporeia are
thought to live for two years or more. Troglobitic
species may live even longer (5 – 6 years) in stable habitats with low but relatively continuous inputs of organic
detritus as food sources. Long-term data demonstrate
19. Introduction to the Subphylum Crustacea
cyclic population oscillations, although control mechanisms for the observed population growth cycles are not
clear. Generally, effects of hydrographic and nutrient
cycles on pelagic primary productivity and related
changes in food quality and quantity on amphipod fecundity and secondary production are well documented
(Hynes and Harper, 1972; Siegfried, 1985; Sarvala,
1986; Johnson and Wiederholm, 1989; Dehdashti and
Blinn, 1991). Isopods inhabiting freshwater and saline
ponds typically have an annual life cycle in contrast to
the longer lifespan of four years for many terrestrial
isopods (Sastry, 1983). Caecidotea recurvata, a cavedwelling isopod, is known to live at least four years (D.
C. Culver, personal communication).
2. Patterns of Reproduction
Generally, Peracarida mate at, or shortly after, the
time of the female molt (Sastry, 1983). In most amphipods and isopods, females produce only a single
brood during the annual life cycle. Seasonal peaks in
annual reproduction occur in Gammarus minus populations in spring habitats but this seasonal pattern in
less distinct in cave populations (Culver et al., 1995).
Some species are known to reproduce continuously and
have multiple broods. For example, the subterranean
isopod Caecidotea tridentata apparently reproduces
throughout the year (Edler and Dodds, 1996). Amphipods, such as Hyalella azteca, can produce multiple
broods during an extended breeding season (Cooper,
1965; Strong, 1972). Adult females can reproduce at
every stage of molting. They carry their eggs in the
marsupium and after fertilization continue to bear their
young until they release the fully developed juveniles
(ranging from a few to more than 20 per brood). Mateguarding is known to occur in some species with timelimited reproductive cycles (e.g., Jormalainen and Shuster, 1999). Some families of amphipods differ in how
mating occurs. The Pontoporeiidae have a pelagic mating system while the Gammaridae generally have a benthic mode (Strong, 1973; Naylor and Adams, 1987;
Bousfield, 1989; Elwood and Dick, 1990; Wen, 1993).
Several types of broad life-history patterns have
been documented for both North American and European species (e.g., Cooper, 1965; Strong, 1972; Gee,
1988; Tadini et al., 1988). In general, the cold-spring
faunas have longer life spans, produce fewer eggs, and
lack marked seasonality in their reproduction compared to warm-water taxa, which only live one year
and produce large numbers of young in the spring. A
slowing down of reproduction during late summer is
reported in some amphipod populations and may be
related to food scarcity or shifts in breeding behavior
controlled by other factors. In Gammarus minus the
onset of amplexus is apparently induced by a rapid
795
decline in temperature (D. C. Culver, personal communication). In widely distributed, warm-water species
such as Hyalella azteca, day length has been ruled out
as a controlling variable for triggering the onset of reproduction (Strong, 1972). For cold-water species such
as Gammarus lacustris, a period of short days and long
nights (typical of winter) is needed to induce reproduction (DeMarch, 1982). In Monoporeia affinis, the
boreo-arctic amphipod, constant illumination inhibits
gonadal development (Sergestrale, 1970).
Peracarid activity patterns on some substrates are
also known to be influenced by the presence of predators (Newman and Waters, 1984; Malmqvist and
Sjostrom, 1987; Holomuzki and Short, 1988, 1990;
Holomuzki and Hoyle, 1990; Holomuzki and Hatchett, 1994; Wooster, 1998). These behavioral changes
may influence life-history patterns in certain habitats.
For example, different populations of the streamdwelling isopod Lirceus fontinalis responded to different risks of predation from banded scuplins (Cottus
carolinae) or streamside salamanders (Ambystoma barbouri) by maturing at larger sizes in those habitats
where gape-limited salamanders were the dominant
predators (Sparkes, 1996a, b). Endoparasites also influence isopod and amphipod behavior and may make
some individuals more vulnerable to predators by increasing their tendency to drift (McCahon et al., 1991;
Hechtel et al., 1993, Bakker et al., 1997; Maynard
et al., 1998). In some cases the infected individuals are
more active during daylight or more visible due to
changed coloration. In these ways the parasite increases
host vulnerability to predators and can enhance transmission of the parasite from the initial crustacean host
to definitive vertebrate hosts such as fishes and birds
that completes the parasite’s life cycle (Bethel and
Homes, 1977; Camp and Huizinga, 1979; Bakker
et al., 1997).
D. Biogeography
The distribution and abundance of any species reflects a large number of variables that may have interacted over very long periods. Climatic changes have
clearly influenced the stability of aquatic habitats and
their availability for colonization by crustaceans for
millions of years. The most recent glacial advances and
retreats were associated with not only major fluctuations in global temperatures, but also with changing
water balances of lakes and river drainage ecosystems.
When much of the earth’s water was frozen in massive
ice sheets, the levels of many lakes and rivers were
much reduced in the temperate zones and throughout
the Neotropics. As discussed later in this section, important refugia from thermal extremes and desiccation
796
A. P. Covich and J. H. Thorp
have persisted in below-ground habitats such as
groundwater-fed caves.
The mechanisms of transport and available routes
of dispersal for a species are primary factors in limiting
access to new habitats and recolonization of old habitats that might have had major climatic disturbances.
Physical and chemical barriers to entry can also limit
range expansions, as may biological barriers such as
the presence of predators or strong competitors already
established in adjacent waters (MacNeil et al., 1997;
1999b). Among the Peracarida, there is a pattern of restricted distribution for many species (sometimes to a
single lake or spring-fed stream) that has interested biologists, geologists, and geographers. A few species are
widespread and seem to tolerate a broad range of ecological parameters. For example, Gammarus lacustris
occurs throughout most of the northern and parts of
the western United States. Hyalella azteca is present
from Canada to South America in the littoral zone of
glacial lakes as well as in small ponds and streams.
Several species of Crangonyx are frequently found in
the southeastern United States as well as in Canada
(Bousfield and Holsinger, 1989). Habitat modification
has changed some amphipod distributions in historic
times, such as the likely effects of dam building and
regulation of flow regimes (e.g., Beckett et al., 1998).
Many species are considered to be relatively rare.
The isopod, Thermosphaeroma theromophilum, has
been declared legally endangered; it is restricted to a
currently protected hot-spring habitat near Socorro,
New Mexico (Bowman, 1981). As discussed below
some cave species are highly restricted in their distributions and some endemic species occur in only a single
cave (Holsinger, 1991; Holsinger, 1994a, b). Recent
reviews provide examples of threats to survival of crustacean species (Elliott, 2000; Ricciardi and Rasmussen,
1999; Culver et al., 2000; Master et al., 2000).
1. Groundwaters
The greatest number of amphipod species are
found in subterranean interstitial habitats, and many
are associated with caves or deep wells (Barr and
Holsinger, 1985; Holsinger, 1993; 1994a). Bousfield
(1983) suggested that more than 1000 hypogean amphipod species will eventually be identified. J. R.
Holsinger (personal communication) completed a
worldwide survey and found that 117 genera have subterranean species; approximately 615 species are fully
adapted to live in groundwaters. There are about 116
species that live in groundwaters in North America,
north of Mexico. Holsinger (1972) estimated that
65 – 70% of the North American amphipod fauna occurs in these groundwater habitats; such species are
known generally as “stygobionts.” He pointed out that
not all of the stygobiont species are restricted to cave
waters per se (true cave-dwelling species are termed
“troglobites”), yet they may have similar morphological specializations associated with cave living (e.g., loss
of eyes and pigmentation). Some cave-dwelling species
are endemic to a single location within one county.
Generally, the geographic locations of obligate cavedwelling species are highly restricted and these subterranean communities appear regionally undersaturated.
Culver et al. (2000) report that over 50% of all the
aquatic obligate cave species and subspecies occur in
only 19 counties (out of 3112) that comprise less than
1% of the land area in the 48 contiguous states. Most
taxa are concentrated in Hays County, Texas or in a
few counties in Florida, Oklahoma, Texas, Virgina, and
West Viriginia. This high concentration of subterranean
taxa in a relatively few locations makes these populations highly vulnerable to habitat destruction. For example, deforestation and dumping of sawdust (into
sink-hole cave from a sawmill operation and leachate
into groundwater from sawdust piles) in southwestern
Virginia led to listing a cave stream isopod (Lirceus usdagalun) as an endangered species (Jacobs, 1991;
Culver et al., 1992). Although many subterranean populations are at risk, only a few have protection through
federal status as rare or endangered.
Groundwaters are insulated from many of the environmental fluctuations that characterize surface waters, but prolonged droughts or massive ice formations
associated with glaciation might eliminate these otherwise “constant” habitats. Until recently, it was thought
that the few subterranean amphipods and isopods
known from glaciated regions represented postglacial
colonization from nonglaciated border regions to the
south. Subglacial refugia are now thought likely to
have persisted during the last glacial episode (known as
the Wisconsin, which ended about 12,000 years ago).
For example, Castleguard Cave in Alberta, Canada, is
currently inhabited by the asellid isopod Salmasellus
steganothrix and the crangonyctid amphipod Stygobromus canadensis. The amphipod is endemic to this single cave and was found about 2 km from the entrance.
Eleven of the 100 species described in the genus Stygobromus are known to occur in various localities north
of the southernmost boundary of the Wisconsin
episode of Pleistocene glaciation (Holsinger et al.,
1983). Thus, the action of glacial scour of surface habitats did act as a barrier to northern distributions of
many species, but others were adapted to subsurface
waters and found refuge.
The fluctuations of sea levels associated with Pleistocene glaciations and earlier Tertiary events also created barriers to distributions of Peracarida. Colonization of coastal streams and subterranean habitats by
19. Introduction to the Subphylum Crustacea
marine-derived species has resulted in a complex
mosaic of biogeographic boundaries. Initially, those organisms that adapted to brackish waters were able to
escape most of their marine-based fish predators and to
obtain abundant food resources in freshwater habitats.
As shallow Cenozoic marine waters receded, the isolated populations of amphipods and isopods developed
distinct adaptations; subsequent speciation produced
many new taxa (Holsinger, 1986; Notenboom, 1991).
In isolated refugia, these species diversified over a 70million year period (possibly much longer) with apparently only slow and limited active dispersal. Because
many of these underground habitats are deep aquifers,
the faunal composition is difficult to sample. The
Edwards Aquifer in south-central Texas has been studied intensively (Holsinger and Longley, 1980), and a
unique ancient fauna of at least 22 species, including
10 amphipod species, is known to live in this stable
groundwater habitat. Abele (1982) concluded that this
artesian well in San Marcos, Texas, contains probably
the richest cave crustacean fauna in North America.
2. Surface Waters
Today, the peracarids occupy many types of surface
waters that range widely in physical and chemical characteristics. Historically, the chemical barrier of dilute
water was a major isolating mechanism (Vernberg and
Vernberg, 1983) in the evolution of crustaceans. In
contrast to the frequently isolated subsurface populations and subpopulations, relatively wider geographic
distributions and frequent gene flow can occur among
surface-dwelling, interbreeding populations in large
lakes and ponds interconnected by rivers. Long-distance transport by individuals adhering to mud carried
by migratory birds, water fowl, and other animals is
considered rare among amphipods but might infrequently provide wider dispersal by those individuals
that do survive short-term drying.
Some species of isopods are reported to move
through subterranean waters and into the surface waters for certain periods (e.g., Minckley, 1961). The importance of hyporheic habitats as spatial and temporal
refugia for many types of benthic crustaceans and insects is now recognized as being of great importance,
especially in previously glaciated regions where massive
unconsolidated rock and sand deposits characterize the
floodplains of large rivers (Stanford and Ward, 1988).
3. Barriers to Dispersal and Population Genetics
Neighboring populations can be genetically isolated from one another by many types of physical barriers. Dispersal of individuals to distant headwaters of
many fast-flowing streams is limited within a drainage
area unless there are subsurface connections among
797
aquifers or many short tributary branches of streams
that connect. Gooch (1989) quantified seven levels of
isolation in his studies of Gammarus minus in the Appalachian Mountains of West Virginia and Pennsylvania. The larger the barrier, the more isolated were the
populations. He has shown that there is a cline of
lower genetic variability along a sequence of northern
populations. These populations have apparently recolonized habitats since the last glacial advance and now
occupy what was once a periglacial zone or perimeter
around the ice sheets (Gooch and Glazier, 1986). The
importance of the geomorphological setting in determining how gammarid populations are actively and
passively distributed is also clear in other studies of
karst populations (e.g., Gooch, 1990; Gooch and
Glazier, 1991; Culver and Fong, 1994).
In addition to the studies on Gammarus mentioned
above, other investigations have focused on Hyalella
and these also provide important new information
about adaptations among distinct populations and subpopulations. For example, in the glacial terrain of
southeastern Michigan’s there is evidence for recent divergence among populations of two ecotypes of
Hyalella azteca (Saussure) that differ in terms of adult
body size (McPeek and Wellborn, 1998). The largebodied adults from different populations found in lakes
interbreed as do the small-bodied adults found in
marshes. In contrast, individual adults of different ecotypes do not interbreed. This type of recent divergence
may have resulted from enhanced mating success favored through sexual selection and mediated by fishpredation selection in the lake populations of different
Hyalella (Wellborn, 1994, 1995, 2000). In Arizona,
there are distinct populations of Hyalella azteca living
in different lake habitats. One group of populations
lives in submerged vegetation while another group
clings to roots of emergent vegetation (Thomas et al.,
1997). Studies have examined genetic differences and
behavioral differences in swimming among different
populations of Hyalella azecta from Oregon to Arizona
(Thomas et al., 1997, 1998) and suggest that xeric
landscapes promote genetic and behavioral divergence
among amphipods, presumably because of the long distances among freshwater habitats. Habitat preferences
and possible segregation of populations likely occur in
other amphipod species living in other structurally complex, stable habitats (e.g., Muller, 1998; Muller et al.,
2000) or habitats that vary temporally and spatially
such as groundwater-fed cave streams and marshes.
Several other studies have considered the importance of habitat structure in the dispersal and population genetics of peracarid crustaceans (Hedgecock
et al., 1982; Meyran and Taberlet, 1998; Meyran et al.,
1997, 1998). The degree of genetic isolation among
798
A. P. Covich and J. H. Thorp
populations distributed within a drainage basin is usually related to topographic complexity and elevational
gradients for many types of slow-moving stream invertebrates (Siegismund and Muller, 1991; Muller, 1998;
Muller et al., 2000). As discussed above, dispersal by
downstream drift and infrequent upstream movements
limits gene flow among subpopulations within river
drainage networks. Long, high ridges dissect the landscape and form effective barriers to the mixing of genes
within and among river networks and associated lake
basins.
4. Comparisons Among Ancient Lakes
Isolation by topographic structure may also be important for benthic crustaceans in deeply dissected lake
basins, as apparently has been the case in the ancient
lakes such as Lake Titicaca in the Andes of Bolivia and
Peru where some 11 species of Hyalella have evolved.
One of the most interesting cases of speciation and
adaptive radiation among freshwater invertebrates is
the high species richness of amphipods in Lake Baikal,
Siberia, the oldest and deepest lake in the world where
nearly 2000 metazoan species occur (Brooks, 1950;
Martens et al., 1994; Yampolsky et al., 1994; Kozhova
and Izmest’va, 1998; Kamaltynov, 1999). Almost 25%
of the total worldwide freshwater amphipod fauna
known to exist today occurs in Baikal, and nearly all of
these species are restricted to this single ancient lake
(Kozhov, 1963; Martens, 1997; Martens and Schon,
1999; Martens and Danielopol, 1999). The progenitors
of the modern fauna can be traced to much earlier
Mesozoic brackish-water basins. Major climatic
changes during the Tertiary are thought to have accelerated the amphipod species radiation within the
Baikal basin during the last 25 – 30 million years (Kamalynov, 1999). Several endemic species also occur
within the Baikal drainage in the Angara and Enisei
Rivers (Kamaltynov, 1992). In some cases, the locations of the outlets and inlets of rivers apparently create barriers for dispersal along the lake shore and result
in genetic separation of different localized populations
(Mashiko et al., 1997). There are at least 250 gammarid species in about 45 genera that have evolved specialized niches in this uniquely deep lake (maximal
depth is over 1700 m). Some are armed with sharp cutaneous spines or ridges, and they range in size from 10
to 80 mm. Coloration is highly varied (red, green, yellow, violet, pink, and brown) with complex patterning,
which is very unusual among freshwater invertebrates.
A phylogenetic analysis of 18 of the endemic genera
suggests that parallel evolution of body armament and
large body size occurred among different genera during
the history of the lake. The first lineage was apparently
benthic and mostly unarmed; the second lineage was
composed of mostly armed taxa and were primarily detritivorous and carnivorous (Sherbakov et al., 1998).
The evolutionary ages of the various taxa are being
studied using molecular techniques such as analysis of
mitrochondrial DNA and allozymes (Mashiko et al.,
1997; Ogarkov et al., 1997; Vainola and Kamaltynov,
1999).
Because of the combination of low water temperatures and very deep circulation, there is sufficient dissolved oxygen in this lake to allow benthic gammarids
to live in several depth zones. Deep-water (1400 m)
forms have evolved different modes of surviving high
hydrostatic pressure (Brauer et al., 1980a; b). Littoral
forms are extremely abundant, reaching densities of
30,000 or more per square meter. One endemic species
of gammarid amphipod, Macrohectopus branickii, is a
dominant pelagic omnivore. Stable isotopic analyses of
N-15 demonstrate that individual consumers change
their mode of feeding to consuming more zooplankton
than phytoplankton as they grow larger (Yoshii et al.,
1999). This relatively large (maximum body length
38 mm) species is a major prey species of several fishes,
especially omul, Coregonus autumnalis migratorius
and pelagic sculpins. In the more productive bays in the
middle basin of Lake Baikal, Macrohectopus forms
dense aggregations and migrates vertically from 250 m
depths to upper waters at night (Melnik et al., 1993).
Using vertical tows of especially designed closing nets
and hydroacoustic equipment Rudstam et al (1992)
were able to determine day and night distributions of
individuals and biomass in Barguzin Bay. They found
that these aggregations were dominated by immature
females (15 mm long) in late summer and contained
some nektobenthic species of amphipods as well. The
migratory patterns and apparent avoidance of fish
predators by Macrohectopus in very similar to Mysis
migrations in Lake Michigan and other North American deep lakes (Lasenby et al., 1986; Rudstam et al.,
1992). Unlike the high species richness of amphipods,
the isopods in Baikal are represented by a single genus,
Asellus, with only five species. Generally, the diversification of crustaceans reflects differences in the response
of isolated populations to local and regional conditions
of long-term habitat stability, plasticity of life-history
characteristics, predator – prey interactions, and access
to limited food resources.
IV. COLLECTING, REARING, AND PREPARATION
FOR IDENTIFICATION
Most amphipods and isopods can be collected by
sweeping a dip net through the littoral zone of lakes
and ponds or in vegetated reaches of streams. Mysids
19. Introduction to the Subphylum Crustacea
require a large diameter plankton net that can be used
for deep vertical tows. Being fast swimmers, mysids can
avoid small nets and are more difficult to collect.
Quantitative sampling of amphipods and isopods has
relied on drift nets and Surber samplers in fast-flowing
streams and Ekman grab sampler in shallow ponds and
lakes. Drift nets can entrap organisms that may be
moving upstream as well as those drifting downstream
(if the net is not positioned off the bottom of the
stream), and, thus, interpretation of studies dealing
with dispersal and migration can be complicated. The
use of cores (e.g., Milstead and Threlkeld, 1986), can
be very effective for intensive study of microhabitats.
Considerable time can be spent in sieving or sorting
specimens; elutriators increase the efficiency of separating specimens from the inorganic and organic matrix
(Magdych, 1981). Baited traps with various mesh sizes
also work well for quantitative sampling of isopods
and gammarids that respond to current-dispersed
chemicals given off by the bait (e.g., Fitzpatrick, 1983;
Allan and Malmqvist, 1989). Sorting of specimens is
relatively rapid, but construction of fine-meshed traps
is necessary. Use of electrofishing equipment may also
be useful in stunning larger decapod crustaceans so
that they can be swept up more effectively with dip
nets. Chemicals have been added to stun or poison animals, but these methods are not very effective and
clearly can be very detrimental to the environment. As
discussed previously, many species are extremely sensitive to even small concentrations of toxic chemicals
and are used to monitor water quality.
Several papers provide details on care and feeding
of laboratory stocks (e.g., Cooper, 1965; Strong, 1972;
DeMarch, 1981a, b; Sutcliffe et al., 1981; Barlocher
et al., 1989). Dead leaves conditioned with nutritious
microbial growths of bacteria and fungi are sufficient
food resources for most species of amphipods and
isopods. These can be supplemented with cultures of
green algae and diatoms or with boiled lettuce or
spinach. As in rearing other taxa, the initial water used
for setting up an aquarium must be well aerated and
allowed to develop an adequate microbial community
before the crustaceans are placed into the tank. Temper-
799
ature fluctuations can be detrimental and some type of
aquarium heater is useful to maintain optimal growing
conditions. Parasites and diseases are, of course, widespread in natural populations (e.g., Pixell Goodrich,
1934; Brownell, 1970; Laberge and McLaughlin,
1989), and brood stock from previously reared laboratory cultures are less likely to suffer from these outbreaks as the complex life histories of many parasites
require a variety of vertebrate and invertebrate hosts to
maintain the life cycle.
Preparation of specimens for study usually requires
rapid fixation and storage of field-collected organisms
in vials of 60 – 70% ethyl alcohol. The alcohol will remove any pigment from the specimens, so careful notes
or photographs will be useful if information on coloration or patterns of body markings are of interest.
The long-term storage of specimens requires checking
to ensure that the alcohol has not evaporated. Addition
of a 5 – 20% solution of glycerin to the vials will protect the specimens from drying out. Glycerin jelly is a
solid at typical room temperatures, and it is widely
used as a permanent mounting medium. Thus, if the
material is first stored in dilute glycerin, it is convenient
to transfer specimens to slides for study and for preparing reference collections. These techniques are described in detail in several texts (e.g., Pennak, 1989;
Peckarsky et al., 1990).
V. CLASSIFICATION OF PERACARIDA
AND BRANCHIURA
The taxonomic key for this chapter is intended to
allow the reader to identify major taxa of amphipods,
isopods, mysids, and Branchiura to the generic level.
This key draws on characteristics used in previously
published studies (Edmondson, 1959; Williams, 1970;
Cressey, 1972; Holsinger, 1972, 1989, 1994b; Fitzpatrick, 1983; Pennak, 1989; Peckarsky et al., 1990);
some of these sources can be used to identify organisms
to species level, others are useful for determining biogeographical distributions (e.g., Barnard and Barnard,
1983; Banarescu, 1990).
A. Taxonomic Key to Major Freshwater Genera of the Orders Mysidacea, Amphipoda, and Isopoda and
the Subclass Branchiura
1a.
Four pairs of legs and two prominent, highly modified suckers on upper ventral surface, specialized ectoparasite on fishes (Fig. 6).....
.................................................................................................................................................Subclass Branchiura, order Arguloida,
family Arguilidae .....................................................................................................................................................................Argulus
1b
More than four pairs of walking legs (pereiopods) prominent second antennae, compound eyes ..................................................... 2
2a(1b)
Five pairs of legs, body flattened laterally (Fig. 7) .................................................................................................order Amphipoda 4
800
A. P. Covich and J. H. Thorp
2b
Six or seven pairs of legs, body flattened dorsoventally .................................................................................................................... 3
3a(2b)
Seven pairs of legs, posterior legs longer than anterior legs, unstalked eyes (Fig.8) .....................................................order Isopoda 8
3b
Six pairs of legs, prominent stalked eyes (Fig. 9) .......................................................................................................order Mysidacea,
family Mysidae ...........................................................................................................................................................................Mysis
4a(2a)
Amphipoda: antenna 1 shorter than antenna 2 (Fig. 7) .................................................................................................................... 5
4b
Antenna 1 longer than antenna 2 (Fig. 7) ......................................................................................................................................... 6
5a (4a)
Accessory flagellum present on antenna I with 3 – 5 segments (Fig. 7), telson deeply cleft but not to base ..........family Pontoporeiidae
.........................................................................................................................................................................................Monoporeia
5b
Accessory flagellum absent on antenna 1, telson entire .................................................family Hyalellidae .............................Hyalella
6a (4b)
Accessory flagellum of antenna 1 with 2 – 7 segments (usually 3 or more), telson typically cleft to base ................family Gammaridae
...........................................................................................................................................................................................Gammarus
6b
Accessory flagellum of antenna 1 with never more than 2 segments, telson usually not deeply cleft but never to the base....................
.....................................................................................................................................................................family Crangonyctidae 7
7a (6b)
Eyes may be present and pigmented, apical margin of telson distinctly cleft........................................................................Crangonyx
7b
Eyes absent, apical margin of telson entire or with shallow cleft ......................................................................................Stygobromus
8a (3a)
Isopoda: margin of head with a pointed median protuberance (carina) between bases of first antennae (Fig. 8), eyes typically present
...........................................................................................................................family Asellidae .............................................Lirceus
8b
Anterior margin of head without carina, eyes present or absent...............................................................................family Asellidae 9
9a (8b)
Abdomen broader than long, up to 16 mm long...................................................................................................................... Asellus
9b
Abdomen not broader than long ....................................................................................................................................... Caecidotea
ACKNOWLEDGMENTS
We greatly appreciate reviews and comments by David C. Culver and Dean W. Blinn as well as help from John R. Holsinger with
several key references. Jennifer L. Borash provided scanning electron
images of crayfish chelae. Rebecca Rudman assisted with a thorough
proof-reading and many authors provided reprints of their studies.
LITERATURE CITED
Abele, L. G. 1982. Biogeography, in: Bliss, D. E., Ed., The biology of
Crustacea. vol. 1. Systematics, the fossil record and biogeography. Academic Press. New York, pp. 242 – 304.
Ache, B. W. 1982. Chemoreception and thermoreception, in: Atwood, H. L., Sandeman, D. C., Eds., The biology of Crustacea.
Vol. 3. Academic Press. New York, pp. 369 – 398.
Adare, K. I., Lasenby, D. C. 1994. Seasonal changes in the total lipid
content of the opossum shrimp, Mysis relicta (Malacostraca,
Mysideacea). Canadian Journal of Fisheries and Aquatic Sciences 51:1935 – 1941.
Allan, J. D., Malmqvist, B. 1989. Diel activity of Gainmarus pulex
(Crustacca) in a South Swedish stream: comparison of drift
catches vs baited traps. Hydrobiologia 179:73 – 80.
Allee, W. C. 1914. The ecological importance of the rheotactic reaction of stream isopods. Biological Bulletin 27:52 – 66.
Allee, W. C. 1929. Studies in animal aggregations: natural aggregations of the isopod, Asellus communis. Ecology 10:14 – 36.
Anderson, R. S., Raasveldt, L. G. 1974. Gammarus predation and the
possible effects of Gammarus and Chaoborus feeding on the zooplankton composition in some small lakes and ponds in western
Canada. Canadian Wildlife Service Occasional Papers 18:1 – 23.
Atema, J. 1988. Distribution of chemical stimuli, in: Atema, J., Ed. Sensory biology of aquatic animals. Springer, New York, pp. 29 – 56.
Bakker, T. C., Mazzi, D., Zala, S. 1997. Parasite-induced changes in
behavior and color make Gammarus pulex more prone to fish
predation. Ecology 78:1098 – 1104.
Banarescu, P. 1990. Zoogeography of fresh waters, Vol. 1. General
distributions and dispersal of freshwater animals. AULA, Wiesbaden. 511 pp.
Banner, A. H. 1953. On a new genus and species of mysid from
southern Louisiana. Tulane Studies in Zoology 1:1 – 8.
Barlocher, F., Kendrick, B. 1973. Fungi and food preferences of Gammarus pseudolimnaeus. Archiv fur Hydrobiologie 72:501 – 516.
Barlocher, F., Kendrick, B. 1975. Assimilation efficiency of Gammarus pseudo! imnaeus (Amphipoda). Oikos 24:295 – 300.
Barlocher, F., Tibbo, P. G., Christie, S. H. 1989. Formation of phenol-protein complexes and their use by two stream invertebrates. Hydrobiologia 173:243 – 249.
Barnard, J. L., Barnard, C. M. 1983. Freshwater Amphipoda of the
world. 1. Evolutionary patterns. 2. Handbook and bibliography. Hayfield Associates, Mt. Vernon, VA.
Barnes, R. D. 1987. Invertebrate Zoology, 5th ed., Saunders, New
York, 893 p.
Barnes, R. D., Harrison, F. W. 1992. Introduction, in: Harrison, F.
W., Humes, A. G., Eds., Microscopic anatomy of invertebrates.
Vol. 9. Crustacea. Wiley – Liss, New York, pp. 1 – 8.
Barr, T. C., Jr. Holsinger, J. R. 1985. Speciation in cave faunas. Annual Review of Ecology and Systematics 16:313 – 337.
Beck, J. T. 1980. Life history relationships between the bopyrid isopod Probopyrus pandalicola and one of its freshwater shrimp
hosts Palaemonetes paludosus. American Midland Naturalist
104:135 – 154.
Beckett, D. C., Lewis, P. A., Green, J. H. 1998. Where have all
the Crangonyx gone? The disappearance of the amphipod
19. Introduction to the Subphylum Crustacea
Crangonyx pseudogracilis, and subsequent appearance of Gammarus nr. fasciatus, in the Ohio River. American Midland Naturalist 139:201 – 209.
Beeton, A. M., Bowers, J. A. 1982. Vertical migration of Mysis relicta Loven. Hydrobiologia 93:53 – 61.
Bethel, W. M., Holmes, J. C. 1977. Altered evasive behavior and responses to light in amphipods harboring acanthaocepthalan cystacanths. Journal of Parasitology 59:945 – 956.
Bohonak, A. J. 1999. Dispersal, gene flow, and population structure.
Quarterly Review of Biology 74:21 – 45.
Blinn, D. W., Davies, R. W. 1990. Concomitant diel vertical migration of a predatory leech and its amphipod prey. Freshwater
Biology 24:401 – 407.
Blinn, D. W., Dehdashti, B., Runck, C., Davies, R. W. 1990. The importance of prey size and density in an endemic predator prey
couple (leech Erpobdella montezuma, amphipod Hyalella montezuma). Journal of Animal Ecology 59:187 – 192.
Blinn, D. W., Grossnickle, N. E., Dehdashti, B. 1987. Diet vertical
migration of a pelagic amphipod in the absence of fish predation. Hydrobiologia 160:165 – 171.
Blinn, D. W., Johnson, D. B. 1982. Filter-feeding of Hyalella montezuma, an unusual behavior for a freshwater amphipod. Freshwater Invertebrate Biology 1:48 – 52.
Blinn, D. W., Pinney, C., Wagner, V. T. 1988. Intraspecific discrimination of amphipod prey by a freshwater leech through
mechanoreception. Canadian Journal of Zoology 66:427 – 430.
Bliss, D. E., Editor-in-Chief, 1982 – 1985. The biology of Crustacea,
Vols. 1 – 10, Academic Press, New York.
Borash, J. L., Moore, P. A. 1997. Ultrastructure, morphology and
distribution of sensory structures on the chelae of the crayfish.
Ohio Journal of Science 97(2): A-11.
Boulton, A. J., Findlay, S., Marmonier, P., Stanley, E. H., Valett,
H. M. 1998. The functional significance of the hyporheic zone
in streams and rivers. Annual Review of Ecology and Systematics 29:59 – 81.
Bousfield, E. L. 1983. An updated phyletic classification and paleohistory of the amphipods, in: Schram, F. R., Ed., Crustacean issues 1: Crustacean phylogeny. Balkema. Rotterdam, pp. 257 –
277.
Bousfield, E. L. 1989. Revised morphological relationships within the
amphipod genera Pontoporeia and Gammaracanthus and the
“glacial relict” significance of their postglacial distributions.
Canadian Journal of Fisheries and Aquatic Sciences 46:
1714 – 1725.
Bousfield, E. L., Holsinger, J. R. 1989. A new crangonyctid amphipod crustacean from hypogean fresh waters of Oregon. Canadian Journal of Zoology 67:963 – 968.
Bowles, D. E., Aziz, K., Knight, C. L. 2000. Macrobrachium (Decapoda: Caridea: Palaemonidae) in the contiguous United
States: a review of the species and an assessmet of threats to
their survival. Journal of Crustacean Biology 20: 158 – 171.
Bowman, T. E. 1981. Thermosphaeroma millerio and T. smithi, new
sphaeromatid isopod crustaceans from hot springs in Chihuahua, Mexico. with a review of the genus. Journal of Crustacean Biology 1:105 – 122.
Bowman, T. E., Abele, L. G. 1982. Classification of the recent Crustacea, in: Abele, L. G., Ed., The biology of Crustacea. Vol. 1.
Systematics. the fossil record, and biogeography. Academic
Press, New York, pp. 1 – 27.
Branstrator, D. K., Cabana, G., Mazumder, A., Rasmussen, J. B.
2000. Measuring life-history ominivory in the opossum shrimp,
Mysis relicta, with stable nitrogen isotopes. Limnology and
Oceanography 45:463 – 467.
Brauer, R. W., Bekman, M. Y., Keyser, J. B., Nesbit, D. L., Shvetzov,
G. N., Wright, S. I. 1980a. Adaptation to high hydrostatic pres-
801
sures of abyssal gammarids from Lake Baikal in eastern Siberia.
Comparative Biochemistry and Physiology 65A:109 – l17.
Brauer, R. W., Bekman, M. Y., Keyser, J. B., Nesbit, D. L., Shvetzov,
S. G., Sidelev, G. N., Wright, S. I. 1980b. Comparative studies
of sodium transport and its relation to hydrostatic pressure
in deep and shallow water gammarid crustaceans from
Lake Baikal. Comparative Biochemistry and Physiology 65A:
119 – 127.
Brooks, J. L. 1950. Speciation in ancient lakes. Quarterly Review of
Biology 25:30 – 60.
Brownell, W. N. 1970. Comparison of Mysis relicta and Pontoporeia
affinis as possible intermediate hosts for the acanthocephalan
Echinorhynchus salmonis. Journal of the Fisheries Research
Board of Canada 27:1864 – 1866.
Buchanan, C., Haney, J. F. 1980. Vertical migrations of zooplankton
in the arctic: a test of the environmental controls, in: Kerfoot,
W. C., Ed., Evolution and ecology of zooplankton communities.
Univ. Press of New England, Hanover, pp. 69 – 79.
Bush, B. M. H., Laverack, M. S. 1982. Mechanoreception, in: Atwood, H. L., Sandeman, D. C., Eds., The biology of Crustacea,
Vol. 3. Academic Press, New York, pp. 399 – 468.
Cabana, G., Tremblay, A., Kalff, J., Rasmussen, J. B. 1994. Pelagic
food chain structure in Ontario lakes: a determinant of mercury
levels in lake trout (Salvelinus namaycush). Canadian Journal of
Fisheries and Aquatic Sciences 51:381 – 389.
Camp, J. W., Huizinga, H. W. 1979. Altered color, behavior and
predation susceptibility of the isopod Asellus intermedius infected with Acanthocephalus dirus. Journal of Parasitology
65:667 – 669.
Capart, A. 1951. Thermobathynella adami, gen. et spec. nov., Anaspidace du Congo Beige. Institut Royal des Sciences et Naturelles
de Belgique, Bulletin 27:1 – 4.
Carpenter, J. H. 1999. Behavior and ecology of Speleonectes epilimnius (Remipedia, Speleonectidae) from surface water of an
anchialine cave on San Salvador Island, Bahamas. Crustaceana
72:979 – 991.
Cavaletto, J. F., Nalepa, T. F., Dermott, R., Gardner, W. S., Quigley,
M. A., Lang, G. A. 1996. Seasonal variation of lipid composition, weight, and length in juvenile Diporeia spp. (Amphipoda)
form Lakes Michigan and Ontario. Canadian Journal of Fisheries and Aquatic Sciences 53:2044 – 2051.
Cheper, N. J. 1980. Thermal tolerance of the isopod Lirceus
brachyurus (Crustacea: Isopoda). American Midland Naturalist
104:312 – 318.
Chess, D. W., Stanford, J. A. 1998. Comparative energetics and lifecycle of the opossum shrimp (Mysis relicta) in native and nonnative environments. Freshwater Biology 40:783 – 794.
Chess, D. W., Stanford, J. A. 1999. Experimental effects of temperature and prey assemblage on growth and lipid accumulation by
Mysis relicta Loven. Hydrobiologia 412:155 – 164.
Chipps, S. R. 1998. Temperature-dependent consumption and gutresidence time in the opossum shrimp Mysis relicta. Journal of
Plankton Research 20:2401 – 2411.
Cole, G. A., Bane, C. A. 1978. Thermosphaeroma subequaluin, n.
gen., n. sp. (Crustacea: Isopoda) from Big Bend National Park,
Texas. Hydrobiologia 59:223 – 228.
Cole, G. A., Watkins, R. L. 1977. HyalelIa montezuma, a new
species (Crustacea: Amphipoda) from Montezuma Well, Arizona. Hydrobiologia 52:175 – 184.
Collart, O. O. 1990. Interaction between the parasite Probopyrus
bithynis (Isopoda, Bopyridae) and one of its hosts, the prawn
Macrobrachium amazonicum (Decapoda, Palaemonidae). Crustaceana 58:258 – 269.
Conlan, K. E. 1994. Amphipod crustaceans and environmental disturbance- a review. Journal of Natural History 28:519 – 554.
802
A. P. Covich and J. H. Thorp
Cook, J. A., Chubb, J. C., Veltkamp, C. J. 1998. Epibionts of Asellus
aquaticus (L.) (Crustacea, Isopoda): a SEM study. Freshwater
Biology 39:423 – 438.
Cooper, S. D., Goldman, C. R. 1982. Environmental factors affecting
predation rates of Mysis relicta. Canadian Journal of Fisheries
and Aquatic Sciences 39:203 – 208.
Cooper, W. E. 1965. Dynamics and production of a natural population of a freshwater amphipod, Hyalella azteca. Ecological
Monographs 35:377 – 394.
Covich, A. P., Fritz, S. C., Lamb, P. J., Marzolf, R. D., Matthews,
W. J., Poiani, K. A., Prepas, E. E., Richman, M. B., Winter, T. C.
1997. Potential effects of climate change on aquatic ecosystems
of the Great Plains of North America.. Hydrological Processes
11:993 – 1021.
Covich, A. P., Palmer, M., Crowl, T. A. 1999. The role of benthic
invertebrate species in freshwater ecosystems. BioScience 49:
119 – 127.
Cressey, R. F. 1972. The genus Argulus (Crustacea: Branchiura) of
the United States. Biota of Freshwater Ecosystems, U.S. Environmental Protection Agency Identification Manual 2:1 – 14.
Crouau, Y. 1989. Feeding mechanisms of the Mysidacea, in: Felgenhauer, B. E., Watling, L., Thistle, A. B., Eds., Functional morphology of feeding and grooming in Crustacea. Balkema, Rotterdam, pp. 153 – 171.
Crowl, T. A., Covich, A. P. 1990. Predator-induced life-history shifts
in a freshwater snail. Science 247:949 – 951.
Culver, D. C., Fong, D. W. 1994. Small-scale and large-scale biogeography of subterranean crustacean faunas of the Virginias. Hydrobiologia 287:3 – 9.
Culver, D. C., Jernigan, R. W., O’Connell, J., Kane, T. C. 1994 The
geometry of natural selection in cave and spring populations of
the amphipod Gammarus minus Say (Crustacea: Amphipoda).
Biological Journal of the Linnean Society 52:49 – 67.
Culver, D. C., Jones, W. K., Holsinger, J. R. 1992. Biological and hydrological investigation of the Cedars, Lee County, Virginia, an
ecologically significant and threatened karst area, in: Stanford,
J. A., Simons, J. J., Eds., Proceedings of the first international
conference on groundwater ecology. American Water Resources
Association, Bethesda, MD, pp. 281 – 290.
Culver, D. C., Kane, T. C., Fong, D. W. 1995. Adaptation and natural selection in caves. The evolution of Gammarus minus. Harvard Univ. Press, Cambridge, MA.
Culver, D. C., Master, L. L., Christman, M. C., Hobbs, H. H., III.
2000. Obligate cave fauna of the 49 contiguous United States.
Conservation Biology 14:386 – 401.
Dadswell, M. J. 1974. Distribution, ecology, and postglacial dispersal
of certain crustaceans and fishes in eastern North America. Publications in Zoology No.11, National Museum of Natural Sciences, National Museums of Canada, Ottawa.
Dahl, E., Emanuelsson, H., von Mecklenburg, C. 1970. Pheromone
transport and reception in an amphipod. Science 170:739 – 740.
Dahl, J. 1998. The impact of vertebrate and invertebrate predators
on a stream benthic community. Oecologia 115:253 – 259.
Dahl, J., Nilsson, P. A., Pettersson, L. B. 1998. Against the flow:
chemical detection of downstream predators in running waters.
Proceedings of the Royal Society of London, series B- Biological
Sciences 265:1339 – 1344.
Danielopol, D. L., Pospisil, P., Rouch, R. 2000. Biodiversity in
groundwater: a large-scale view. Trends in Ecology and Evolution 15:223 – 224.
Dehdashti, B., Blinn, D. W. 1991. Population dynamics and production of the pelagic amphipod Hyalella montezuma in a thermally constant system. Freshwater Biology 25:131 – 141.
Delong, M. D., Summers, R. B., Thorp, J. H. 1993. Influence of
food type on the growth of a riverine amphipod, Gammarus
fasciatus. Canadian Journal of Fisheries and Aquatic Sciences
50: 1891 – 1896.
DeMarch, B. G. E. 1981a. Hyalella azteca (Saussure), in: Lawrence,
S. G., Ed., Manual for the culture of selected freshwater invertebrates. Canadian Special Publication in Fisheries and Aquatic
Sciences No. 54, pp. 61 – 77.
DeMarch, B. G. E. 1981b. Gammarus lacustris lacustris G. 0. Sars,
in: Lawrence, S. G., Ed., Manual for the culture of selected
freshwater invertebrates. Canadian Special Publication in Fisheries and Aquatic Sciences No. 54, pp. 79 – 94.
DeMarch, B. G. E. 1982. Decreased day length and light intensity as
factors inducing reproduction in Gammarus lacustris lacustris
Sars. Canadian Journal of Zoology 60:2962 – 2965.
Dermott, R. M., Corning, K. 1988. Seasonal ingestion rates of Pontoporeia hoyi (Amphipoda) in Lake Ontario. Canadian Journal
of Fisheries and Aquatic Sciences 45:1886 – 1895.
Dick, J. T. A. 1995. The cannibalistic behaviour of two Gammarus
species (Crustacea: Amphipoda). Journal of Zoology (London)
236:697 – 706.
Dick, J. T. A., Elwood, R. W., Montgomery, W. I. 1995. The behavioural basis of a species replacement: differential aggression and
predation between the introduced Gammarus pulex and the native G. duebeni celticus (Amphipoda). Behavioral Ecology and
Sociobiology 37:393 – 398.
Dodds, W. K. 1990. Hydrodynamic constraints on evolution of chemically mediated interactions between aquatic organisms in unidirectional flows. Journal of Chemical Ecology 16:1417 – 1430.
Dodson, S. I., Crowl, T. A., Peckarsky, B. L., Kats, L. B., Covich,
A. P., Culp, J. M. 1994. Non-visual communication in freshwater benthos: an overview. Journal of the North American Benthological Society 13:268 – 282.
Donner, K. O., Lindstrom, A., Lindstrom, M. 1987. Seasonal variation in the vertical migration of Pontoporeia affinis (Crustacea,
Amphipoda). Annales Zoologica Fennica 24:305 – 313.
Dormaar, K. A., Corey, S. 1978. Some aspects of osmoreculation in
Mysis relicta Loven (Mysidacea). Crustaceana 34:90 – 93.
Drost, C. A., Blinn, D. W. 1997. Invertebrate community of Roaring
Springs Cave, Grand Canyon National Park, Arizona. Southwestern Naturalist 42:497 – 500.
Dunham, P. 1978. Sex pheromones in Crustacea.. Biological Review
53:555 – 583.
Edler, C., Dodds, W. K. 1996. The ecology of a subterranean isopod,
Caecidotea tridentata. Freshwater Biology 35:249 – 259.
Edmondson, W. T. Ed. 1959. Freshwater biology, 2nd ed., Wiley,
New York.
Elliott, W. R. 1999. Conservation of the North American cave and
karst biota, in: Wilken, H., Culver, D. C., Humphreys, W., Eds.,
Subterranean ecosystems. Elsevier, Oxford, UK, pp. 671 – 695.
Ellis, R. J. 1971. Notes on the biology of the isopod Asellus tomalensis Harford in an intermittent pond. Transactions of the American Microscopical Society 90:51 – 61.
Elwood, R. W., Dick, J. T. A. 1990. The amorous Gammarus: The
relationship between precopula duration and size assortative
mating in G. pulex. Animal Behaviour 39:828 – 833.
Emerson, M. J., Schram, F. R. 1990. The origin of crustacean biramous appendages and the evolution of Arthropoda. Science
250:667 – 669.
Enright, J. T. 1977. Diurnal vertical migration: adaptive significance
and timing. Part I. Selective advantage: A metabolic model.
Limnology and Oceanography 22:856 – 872.
Enright, J. T. 1979. The why and when of up and down. Limnology
and Oceanography 24:788 – 791.
Evans, M. S., Bathelt, R. W., Rice, C. P. 1982. Polychlorinated
biphenyls and other toxicants in Mysis relicta. Hydrobiologia
93:205 – 215.
19. Introduction to the Subphylum Crustacea
Felgenhauer, B. E., Abele, L. G., Felder, D. L. 1992. Remipedia, in:
Harrison, F. W., Humes, A. G., Eds., Microscopic anatomy of
invertebrates, Vol. 9 Crustacea. Wiley – Liss, New York, pp.
225 – 247.
Findlay, S., Meyer, J. L., Smith, J. P. 1984. Significance of bacterial
biomass in the nutrition of a freshwater isopod (Lirceus sp.).
Oecologia 63:38 – 42.
Findlay, S., Meyer, J. L., Smith, P. J. 1986. Contribution of fungal
biomass to the diet of a freshwater isopod (Lirceus sp.). Freshwater Biology 16:377 – 385.
Fitzpatrick, J. F., Jr. 1983. How to know the freshwater Crustacea.
Brown. Dubuque, IA. 227 p.
Folt, C. L., Rybock, J. T., Goldman, C. R. 1982. The effect of prey
composition and abundance on the predation rate and selectivity of Mysis relicta. Hydrobiologia 93:133 – 143.
Fong. D. W. 1989. Morphological evolution of the amphipod Gammarus minus in caves-quantitative genetic analysis. American
Midland Naturalist 121:361 – 378.
Fong, D. W., Culver, D. C. 1994. Fine-scale biographic differences in
the crustacean fauna of a cave system in West Virginia, USA.
Hydrobiologia 287:29 – 37.
France, R. L. 1992. The North American latitudinal gradient in
species richness and geographical range of freshwater crayfish
and amphipods. American Naturalist 139:342 – 354.
France, R. L. 1993. Production and turnover of Hyalella azteca in
central Ontario, Canada compared with other regions. Freshwater Biology 30:343 – 349.
Friesen, J. A., Mann, K. H., Novitsky, J. A. 1986. Mysis digests cellulose in the absence of a gut micro-flora. Canadian Journal of
Zoology 64:442 – 446.
Galat, D. L., Coleman, M., Robinson, R. 1988. Experimental effects
of elevated salinity on three benthic invertebrates in Pyramid
Lake, Nevada. Hydrobiologia 158:133 – 144.
Gee, J. H. R. 1988. Population dynamics and morphometrics of
Gammarus pulex L.: evidence of seasonal food limitation in a
freshwater detritivore. Freshwater Biology 19:333 – 343.
Glazier, D. S. 1999. Variation in offspring investment within and
among populations of Gammarus minus Say (Crustacea: Amphipoda) in ten mid-Appalachian springs (USA). Archiv fur Hydrobiologie 146:257 – 283.
Glazier, D. S., Horne, M. T., Lehman, M. E. 1992. Abundance, body
composition and reproductive output of Gammarus minus
(Crustacea: Amphipoda) in ten cold springs differing in pH and
ionic content. Freshwater Biology 28:149 – 163.
Glazier, D. S., Sparks, B. L. 1997. Energetics of amphipods in ionpoor waters: Stress resistance is not invariably linked to low
metabolic rates. Functional Ecology 11:126 – 128.
Gliwicz, M. Z. 1986. Predation and the evolution of vertical migration in zooplankton. Nature 320:746 – 748.
Goddard, J. S. 1988. Food and feeding, in: Holdich, D. M., Lowery,
R. S., Eds., Freshwater crayfish. Biology, management and exploitation. Timber Press. Portland. OR, pp. 145 – 166.
Goedkoop, W., Johnson, R. K. 1994. Exploitation of sediment bacterial carbon by juveniles of the amphipod Monoporeia affinis.
Freshwater Biology 32:553 – 563.
Gooch, J. L. 1989. Genetic differentiation in relation to stream distance in Gammarus minus (Crustacea, Amphipoda) in Appalachian watersheds. Archiv fur Hydrobiologie 114:505 – 519.
Gooch, J. L. 1990. Spatial genetic patterns in relation to regional history and structure — Gammarus minus (Amphipoda) in Appalachian
watersheds.
American
Midland
Naturalist
124:93 – 104.
Gooch, J. L., Glazier, D. S. 1986. Levels of heterozvgosity in the amphipod Gammarus minus in an area affected by Pleistocene
glaciation. American Midland Naturalist 116:57 – 63.
803
Gooch, J. L., Glazier, D. S. 1991. Temporal and spatial patterns in
mid-Appalachian springs. Memoirs of the Entomological Society of Canada 155:29 – 49.
Graca, M. A. S., Maltby, L., Calow, P. 1994a. Comparative ecology
of Gammarus pulex (L.) and Asellus aquaticus (L.) I: population dynamics and microdistribution. Hydrobiologia 281:155 –
162.
Graca, M. A. S., Maltby, L., Calow, P. 1994b. Comparative ecology
of Gammarus pulex (L.) and Asellus aquaticus (L.) II: Fungal
preferences. Hydrobiologia 281:163 – 170.
Greenaway, P. 1985. Calcium balance and molting in the Crustacea.
Biological Reviews of the Cambridge Philosophical Society
60:425 – 454.
Haden, A., Blinn, D. W., Shannon, J. P., Wilson, K. P. 1999. Interference competition between the net-building caddisfly Ceratopsyche oslari and the amphipod Gammarus lacustris. Journal of
Freshwater Ecology 14:277 – 280.
Hairston, N. G., Bohonak, A. J. 1998. Copepod reproductive strategies: life-history theory, phylogenetic pattern and invasion of inland waters. Journal of Marine Systems 15:23 – 34.
Hairston, N. G., Caceres, C. E. 1996. Distribution of crustacean diapause: micro- and macroevolutionary pattern and process. Hydrobiologia 320:27 – 44.
Hammer, U. T., Sheard, J. S., Kranabetter, J. 1990. Distribution and
abundance of littoral benthic fauna in Canadian prairie saline
lakes. Hydrobiologia 197:173 – 192.
Hartnoll, R. G. 1982. Growth, in: Abele, L. G., Ed., The biology of
Crustacea, Vol. 2. Academic Press, New York, pp. 111 – 196.
Hechtel, L. J., Johnson, C. L., Juliano, S. A. 1993. Modification of
antipredator behavior of Caecidotea intermedius by its parasite
Acanthocephalus dirus. Ecology 74:710 – 713.
Hedgecock, D., Tracey, M. L., Nelson, K. 1982. Genetics, in: Abele,
L. G., Ed., The biology of Crustacea, Vol. 2. Academic Press,
New York, pp. 284 – 403.
Helluy, S., Holmes, J. C. 1990. Serotonin, octopamine, and the clinging behavior induced by the parasite Polymorphus paradoxus
(Acanthocephala) in Gammarus lacustris (Crustacea). Canadian
Journal of Zoology 68:1214 – 1220.
Henry, K. S., Danielopol, D. L. 1998. Oxygen dependent habitat selection in surface and hyporheic environments by Gammarus
roeseli Gervais (Crustacea, Amphipoda): experimental evidence.
Hydrobiologia 390:51 – 60.
Hervant, F., Mathieu, J., Culver, D. C. 1999. Comparative responses
to severe hypoxia and subsequent recovery in closely related
amphipod populations (Gammarus minus) from cave and surface habitats. Hydrobiologia 392:197 – 204.
Hill, C., Quigley, M. A., Cavaletto, J. F., Gordon, W. 1992. Seasonal
changes in lipid content and composition in the benthic amphipods Monoporeia affinis and Pontoporeia femorata. Limnology and Oceanography 37:1280 – 1289.
Hogg, I. D., Williams, D. D. 1996. Response of stream invertebrates
to a global-warming thermal regime: An ecosystem-level manipulation. Ecology 77:395 – 407.
Hogg, I. D., Eadie, J. M., de Lafontaine, Y. 1998a. Atmospheric
change and the diversity of aquatic invertebrates: Are we missing the boat? Environmental Monitoring and Assessment
49:291 – 301.
Hogg, I. D., Larose, C., de Lafontaine, Y., Doe, K. G. 1998b. Genetic
evidence for a Hyalella species complex within the Great Lakes
St. Lawrence River drainage basin: implication for ecotoxicology and conservation biology. Canadian Journal of Zoology
76:1134 – 1140.
Holomuzki, J. R., Hatchett, L. A. 1994. Predator avoidance costs
and habituation to fish chemicals by a stream isopod. Freshwater Biology 32:585 – 592.
804
A. P. Covich and J. H. Thorp
Holomuzki, J. R., Hoyle, J. D. 1990. Effect of predatory fish presence on habitat use and diel movement of the stream amphipod,
Gammarus minus. Freshwater Biology 24:509 – 517.
Holomuzki, J. R., Short, T. M. 1988. Habitat use and fish avoidance
behaviors by the stream-dwelling isopod Lirceus fontinalis.
Oikos 52:79 – 86.
Holomuzki, J. R., Short, T. M. 1990. Ontogenetic shifts in habitat
use and activity in a stream-dwelling isopod. Holarctic Ecology
13:300 – 307.
Holsinger, J. R. 1972. The freshwater amphipod crustaceans (Gammaridae) of North America. Biota of Freshwater Ecosystems,
Identification Manual 5, U.S. Environmental Protection Agency.
Holsinger, J. R. 1986. Zoogeographic patterns of North American
subterranean amphipod crustaceans, in: Gore, R. H., Heck, K.
L., Eds., Crustacean biogeography. Balkema, Rotterdam, pp.
85 – 106.
Holsinger, J. R. 1988. Troglobites: the evolution of cave-dwelling organisms. American Scientist 76:146 – 153.
Holsinger, J. R. 1989. Allocrangonyctidae and Pseudocrangonyctidae, two new families of Holarctic subterranean amphipod
crustaceans (Gammaridae), with comments on their phylogenetic and zoogeographic relationships. Proceedings of the Biological Society of Washington 102:947 – 959.
Holsinger, J. R. 1991. Species accounts: freshwater amphipods and
freshwater isopods, in Hoffman, R. L., Ed., Arthropods in Virginia’s endangered species. McDonald and Woodward, Blacksburg, VA, pp. 180 – 199.
Holsinger, J. R. 1993. Biodiversity of subterranean amphipod crustaceans: global patterns and zoogeographic implications. Journal of Natural History 27:821 – 835.
Holsinger, J. R. 1994a. Pattern and process in the biogeography of
subterranean amphipods. Hydrobiologia 287:131 – 145.
Holsinger, J. R. 1994b. Amphipoda, in: Juberthie, C., Decu, V., Eds.,
Encyclopaedia Biospeologica, tome I, Societe de Biospeologie,
Moulis-Bucarest, pp. 147 – 163.
Holsinger, J. R., Hubbard, D. A., Jr., Bowman, T. E. 1994. Biogeographic and ecological implications of newly discovered populations of the stygobiont isopod crustacean Antrolana lira Bowman (Cirolanidae). Journal of Natural History 28:1047 – 1058.
Holsinger, J. R., Longley, G. 1980. The subterranean amphipod crustacean fauna of an artesian well in Texas. Smithsonian Contributions to Zoology 308:1 – 62.
Holsinger, J. R., Mort, J. S., Reckles, A. D. 1983. The subterranean
crustacean fauna of Castleguard Cave, Columbia Icefields, Alberta, Canada, and its zoogeographical significance. Arctic and
Alpine Research 15:543 – 549.
Hubricht, L., Mackin, J. G. 1949. The American isopods of the
genus Lirceus (Asellota, Asellidae). American Midland Naturalist 42:334 – 349.
Hutchinson, G. E. 1967. A treatise on limnology, Vol. 2. Wiley, New
York. 1115 pp.
Hutchinson, G. E. 1993. A treatise on limnology, Vol. 4. Wiley, New
York. 944 pp.
Hynes, H. B. N., Harper, F. 1972. The life histories of Gammarus lacustris and G. pseudolimnacus in southern Ontario. Crustaceana (Supplement) 3:329 – 341.
Jacobs, J. 1991. Endangered and threatened wildlife and plants; proposal to list the Lee County cave isopod (Lirceus usdagalun) as
an endangered species. Federal Register 56 (221):58026 – 58029.
Johannsson, O. 1992. Life history and productivity of Mysis relicta
in Lake Ontario. Journal of Great Lakes Research 18:154 – 168.
Johannsson, O. E. 1995. Response of Mysis relicta population dynamics and productivity to spatial and seasonal gradients in
Lake Ontario. Canadian Journal of Fisheries and Aquatic Sciences 52:1509 – 1522.
Johannsson, O. E., Rudstam, L. G., Lasenby, D. C. 1994. Mysis
relicta assessment of metalimnetic feeding and implications
for competition with fish in lakes Ontario and Michigan.
Canadian Journal of Fisheries and Aquatic Sciences 51:
2591 – 2602.
Johnson, R. K., Wiederholm, T. 1989. Long-term growth oscillations
of Pontoporeia affinis Lindstrom (Crustacea: Amphipoda) in
Lake Malaren. Hydrobiologia 175:183 – 194.
Johnson, R. K., Wiederholm, T. 1992. Pelagic – benthic coupling —
the importance of diatom interannual variability for population
oscillations of the amphipod Monoporeia affinis (Lindstrom).
Limnology and Oceanography 37:1596 – 1607.
Jones, R., Culver, D. C. 1989. Evidence for selection on sensory
structures in a cave population of Gammarus minus (Amphipoda). Evolution 43:688 – 693.
Jormalainen, V., Shuster, S. M. 1997. Microhabitat segregation and
cannibalism in an endangered freshwater isopod, Thermosphaeroma thermophilium. Oecologia 111:271 – 279.
Jormalainen, V., Shuster, S. M. 1999. Female reproductive cycle and
sexual conflict over precopulatory mate-guarding in Thermosphaeroma (Crustacea, Isopoda). Ethology 105:233 – 246.
Jormalainen, V., Shuster, S. M., Wildey, H. C. 1999. Reproductive
anatomy, precopulatory mate guarding, and paternity in the Socorro isopod, Thermosphaeroma thermophilum, Marine and
Freshwater Behaviour and Physiology 32:39 – 56.
Juday, C., Birge, E. A. 1927. Pontoporeia and Alysis in Wisconsin
lakes. Ecology 7:445 – 452.
Kamaltynov, R. M. 1992. On the present state of systematics of the
Lake Baikal amphipods (Crustacea, Amphipoda). Hydrobiological Journal 26:82 – 92.
Kamaltynov, R. M. 1999. On the evolution of Lake Baikal amphipods. Crustaceana 72:921 – 931.
Kane, T. C., Culver, D. C., Jones, R. T. 1992. Genetic structure of
morphologically differentiated populations of the amphipod
Gammarus minus. Evolution 46:272 – 278.
Kane, T. C., Mathieu, J., Culver, D. C. 1995. Biotic fluxes and gene
flowages, in: Gibert, J., Danielopol, D. L., Stanford, J. A, Eds.,
Groundwater ecology. Academic Press, San Diego, CA, pp.
245 – 270.
Knapp, S. M., Fong, D. W. 1999. Estimates of population size of Stygobromus emarginatus (Amphipoda: Crangonyctidae) in a
headwater stream in Organ Cave, West Virginia. Journal of
Cave and Karst Studies 61:3 – 6.
Koop, J. H. E., Grieshaber, M. K. 2000. The role of ion regulation
in the control of the distribution of Gammarus tigrinus (Sexton)
in salt-polluted rivers. Journal of Comparative Physiology B.
Biochemical Systematic and Environmental Physiology 170:
75 – 83.
Kozhov, N. I. 1963. Lake Baikal and its life, Dr. W. Junk, The
Hague, 344 p.
Kozhova, O. M., Izmest’va, L. R. 1998. Lake Baikal: evolution and
diversity, Backhuys Publishers, Leiden, The Netherlands.
Laberge, R. J. A., McLaughlin, J. D. 1989. Hyalella azteca (Amphipoda) as an intermediate host of the nematode Streptocara
crassicauda. Canadian Journal of Zoology 67:2335 – 2340.
Lamarre, E., Cochran, P. A. 1992. Lack of host species slection by
the exotic parasitic crustacean, Argulus japonicus. Journal of
Freshwaer Ecology 7:77 – 80.
Landrum, P. F., Nalepa, T. F. 1998. A review of the factors affecting
the ecotoxicology of Diporeia spp. Journal of Great Lakes Research 24:889 – 904.
Lasenby, D. C., Northcote, T. G., Furst, M. 1986. Theory, practice
and effects of Mysis relicta introduction to North American and
Scandinavian lakes. Canadian Journal of Fisheries and Aquatic
Sciences 43:1277 – 1284.
19. Introduction to the Subphylum Crustacea
Lee, C. E., Bell, M. A. 1999. Causes and consequences of recent
freshwater invasions by saltwater animals. Trends in Ecology
and Evoutlion 14:284 – 288.
Lester, G. T., Clark, W. H. Occurrence of Corophium spinicore
Stimpson 1857 (Amphipoda: Corophiidae) in Idaho, U.S.A.
Western North American Naturalist (in press).
Lopez, G., Elmgren, R. 1989. Feeding depths and organic absorption
for the deposit-feeding benthic amphipods Pontoporeia affinis
and Pontoporcia femorata. Limnology and Oceanography
34:982 – 991.
MacNeil, C., Dick, J. T., Elwood, R. W. 1997. The trophic ecology of
freshwater Gammarus spp. (Crustacea: Amphipoda): Problems
and perspectives concerning the functional feeding group concept. Biological Reviews of the Cambridge Philosophical Society
72:349 – 364.
MacNeil, C., Dick, J. T. A., Elwood, R. W. 1999. The dynamics of
predation on Gammarus spp. (Crustacea: Amphipoda). Biological Reviews of the Cambridge Philosophical Society
74:375 – 395.
MacNeil, C., Elwood, R. W., Dick, J. T. A. 1999a. Predator-prey interactions between brown trout Salmo trutta and native and introduced amphipods: their implications for fish diets. Ecography 22:686 – 696.
MacNeil, C., Elwood, R. W., Dick, J. T. A. 1999b. Differential microdistributions and interspecific interactions in coexisting
Gammarus and Crangonyx amphipods. Ecography 22:415 –
423.
Magdych, W. P. 1981. An efficient, inexpensive elutriator design for
separating benthos from sediment samples. Hydrobiologia
85:157 – 159.
Maguire, B. 1965. Monodella texana, an extension of the range of
the crustacean order Thermosbaenacea to the western hemisphere. Crustaceana 9:149 – 154.
Malard, F., Hervant, F. 1999. Oxygen supply and the adaptations of
animals in groundwater. Freshwater Biology 41:1 – 30.
Malmqvist, B., Sjostrom, P. 1987. Stream drift as a consequence of
disturbance by invertebrate predators. Field and laboratory experiments. Oecologia 74:396 – 403.
Maltby, L. 1991. Pollution as a probe of life-history adaptations in
Asellus aquaticus (Isopoda). Oikos 61:11 – 18.
Maltby, L. 1995. Sensitivity of the crustaceans Gammarus pulex (L.)
and Asellus aquaticus (L.) to short-term exposure to hypoxia
and unionzed ammonia- observations and possible mechanisms.
Water Research 29:781 – 787.
Mantel, L. H., Farmer, L. L. 1983. Osmotic and ionic regulation, in:
Mantel, L. H., Ed., The biology of Crustacea, Vol. 5 Academic
Press, New York, pp. 53 – 161.
Marchant, R., Hynes, H. B. N. 1981a. The distribution and production of Gammarus pseudolimnaeus (Crustacea: Amphipoda)
along a reach of the Credit River, Ontario. Freshwater Biology
11:169 – 182.
Marchant, R., Hynes, H. B. N. 1981b. Field estimates of feeding rate
for Gammarus pseudolimnaeus (Crustacea: Amphipoda) in the
Credit River, Ontario. Freshwater Biology 11:27 – 36.
Margolis, L., Kabata, Z., Eds. 1988. Guide to the parasites of fishes
of Canada, Part II. Crustacea. Canadian Special Publication of
Fisheries and Aquatic Sciences 101. Department of Fisheries
and Oceans, Ottawa. 184 p.
Martens, K. 1997. Speciation in ancient lakes. Trends in Ecology and
Evolution 12:177 – 182.
Martens, K., Coulter, G., Goddeeris, B. 1994. Speciation in ancient
lakes 40 years after Brooks. Advances in Limnology 44:75 – 96.
Martens, K., Danielopol, D. L. 1999. Concluding remarks- age and
origin of crustacean deiversity in “extreme” environments.
Crustaceana 72:1031 – 1037.
805
Martens, K., Schon, I. 1999. Crustacean biodiversity in ancient lakes:
a review. Crustaceana 72:899 – 910.
Martinez, P. J., Bergersen, E. P. 1989. Proposed biological management of Mysis relicta in Colorado lakes and reservoirs. North
American Journal of Fisheries Management 9:1 – 11.
Marzolf, G. R. 1965. Substrate relations of the burrowing amphipod
Pontoporeia affinis in Lake Michigan. Ecology 46:579 – 592.
Mashiko, K., Kamaltynov, R. M., Shervakov, D. Y., Morino, H.
1997. Genetic separation of gammarid (Eulimnogammarus cyaneus) populations by localized topographic changes in ancient
Lake Baikal. Archiv fur Hydrobiologie 139:379 – 387.
Master, L. L., Stein, B. A., Kutner, L. S., Hammerson, G. A. 2000.
Vanishing assets: the conservation status of U.S. species, in:
Stein, B. A., Kutner, L. S., Adams, J. S., Eds., Precious heritage:
The status of biodiversity in the United States. Oxford Univ.
Press, New York, pp. 93 – 118.
Maynard, B. J., Wellnitz, T. A., Wright, W. G., Dezfull, B. S. 1998.
Parasite-altered behavior in a crustacean intermediate host: field
and laboratory studies. Journal of Parasitology 84:1102 – 1106.
McCahon, C. P., Maund, S. J., Poulton, M. J. 1991. The effect of the
acanthocephalan parasite (Pomphorhynchus laevis) on the drift
of its intermediate host (Gammarus pulex). Freshwater Biology
25:507 – 513.
McDonald, M. E., Crowder, L. B., Brandt, S. B. 1990. Changes in
Mysis and Pontoporcia populations in southeastern Lake
Michigan: a response to shifts in the fish community. Limnology
and Oceanography 35:220 – 227.
McLaughlin, P. A. 1980. Comparative morphology of recent Crustacea. Freeman. San Francisco. 177 p.
McLaughlin, P. A. 1982. Comparative morphology of crustacean appendages, in: Abele, L. G., Ed., The biology of Crustacea, Vol.
2. Academic Press, New York, pp. 197 – 256.
McLoughlin, N. J., Blinn, D. W., Davies, R. W. 1999. An energetic
evaluation of a predator-prey (leech-amphipod) couple in Montezuma Well, Arizona, USA. Functional Ecology 13:45 – 50.
McPeek, M. A., Wellborn, G. A. 1998. Genetic variation and reproductive isolation among phenotypically divergent amphipod
populations. Limnology and Oceanography 43:1162 – 1169.
Melnik, N. G., Timoshkin, O. A., Sideleva, V. G., Pushkin, S. V.,
Mamylov, V. S. 1993. Hydroacoustic measurement of the density of the Baikal macrozooplankter Macrohectopus branickii
(Dyb.) (Amphipods, Gammaridae). Limnology and Oceanography 38:425 – 434.
Meyran, J-C. 1997. Impact of water calcium on the phenotypic diversity of alpine populations of Gammarus fossarum. Ecology
78:1579 – 1587.
Meyran, J-C. 1998. Ecophysiological diversity of alpine populations
of Gammarus lacustris in relation to environmental calcium.
Freshwater Biology 39:41 – 47.
Meyran, J-C., Gielly, L., Taberlet, P. 1998. Environmental calcium,
and mitochondrial DNA polymorphism among local populations of Gammarus fossarum (Crustacea, Amphipoda). Molecular Ecology 7:1391 – 1400.
Meyran, J-C., Monnerot, M., Taberlet, P. 1997. Taxonomic status
and phylogenetic relationships of some species of the genus
Gammarus (Crustacea, Amphipoda) deduced from mitochrondrial DNA sequences. Molecular Phylogenetics and Evolution
8:1 – 10.
Meyran, J-C., Taberlet, P. 1998. Mitochondrial DNA polymorphism
among alpine populations of Gammarus lacustris (Crustacea,
Amphipoda). Freshwater Biology 39:259 – 265.
Milstead, B., Threlkeld, S. T. 1986. An experimental analysis of
darter predation on Hyalella azteca using semipermeable enclosures. Journal of the North American Benthological Society
5:311 – 318.
806
A. P. Covich and J. H. Thorp
Minckley, W. L. 1961. Occurrence of subterranean isopods in the
epigean environment. American Midland Naturalist 63:452 –
455.
Monk, D. C. 1977. The digestion of cellulose and other dietary components, and pH of the gut in the amphipod Gammarus pulex
(L.). Freshwater Biology 7:431 – 440.
Morgan, M. D., Threlkeld, S. T., Goldman, C. R. 1978. Impact of
the introduction of kokanee (Oncorhyncus nerka) and opossum
shrimp (Mysis relicta) on a subalpine lake. Journal of the Fisheries Research Board of Canada 35:1572 – 1579.
Mosslacher, F. 1998. Subsurface dwelling crustaceans as indicators of
hydrological conditions, oxygen concentrations, and sediment
structure in an alluvial aquifer. International Review of Hydrobiology 83:349 – 364.
Muller, J. 1998. Genetic population structure of two cryptic Gammarus fossarum types across a contact zone. Journal of Evolutionary Biology 11:79 – 101.
Muller, J., Partsch, E., Link, A. 2000. Differentiation in morphology
and habitat partitioning of genetically characterized Gammarus
fossarum forms (Amphipoda) across a contact zone. Biological
Journal of the Linnean Society 69:41 – 53.
Mullis, R. M., Revitt, D. M., Shutes, R. E. 1996. A statistical approach for the assessment of the toxic influences on Gammarus
pulex (Amphipoda) and Asellus aquaticus (Isopoda) exposed to
urban aquatic discharges. Water Research 30:1237 – 1243.
Murtaugh, P. A. 1989. Fecundity of Neomysis mercedis Holmes in
Lake Washington (Mysidacea). Crustaceana 57:194 – 200.
Naylor, C., Adams, J. 1987. Sexual dimorphism, drag constraints
and male performance in Gammarus duebeni (Amphipoda).
Oikos 48:23 – 27.
Naylor, C., Pindar, L., Calow, P. 1990. Inter and intraspecific variation in sensitivity to toxins: the effects of acidity and zinc on the
freshwater crustaceans Asellus aquaticus (L.) and Gammarus
pulex (L.). Water Research 24:757 – 762.
Nero, R. W., Schindler, D. W. 1983. Decline of Mysis relicta during
acidification of Lake 223. Canadian Journal of Fisheries and
Aquatic Sciences 40:1905 – 1911.
Newman, R. M. 1991. Herbivory and detritivory on freshwater
macrophytes by invertebrates: A review. Journal of the North
American Benthological Society 10:89 – 114.
Newman, R. M., Hanscom, Z., Kerfoot, W. C. 1992. The watercress
glucosinolate-myrosinase system: a feeding deterrent to caddisflies, snails and amphipods. Oecologia 92:1 – 7.
Newman, R. M., Kerfoot, W. C., Hanscom, Z., III. 1990. Watercress
and amphipods: potential chemical defense in a spring stream
macrophyte. Journal of Chemical Ecology 16:245 – 259.
Newman, R. M., Perry, J. A., Tam, E., Crawford, R. L. 1987. Effects
of chronic chlorine exposure on litter processing in outdoor experimental streams. Freshwater Biology 18:415 – 428.
Newman, R. M., Waters, T. F. 1984. Size-selective predation on
Gammarus pseudolimneaus by trout and sculpins. Ecology
65:1535 – 1545.
Nolan, D. T., van der Salm, A. L., Bonga, S. E. W. 2000. The hostparasite relationship between the rainbow trout (Oncorhynchus
mykiss) and the ectoparasite Argulus foliaceus (Crustacea:
Branchiura): epithelial mucous cell response, corticol and factors which may influence parsite establishment. Contributions
to Zoology 69:57 – 63.
Notenboom, 1991. Marine regression and the evolution of groundwater dwelling amphipods (Crustacea). Journal of Biogeography 18:437 – 454.
Notenboom, J., Plenet, S., Turquin, M. J. 1994. Groundwater contamination and its impact on groundwater animals and ecosystems,
in: Gibert, J., Danielopol, D. L., Stanford, J. A., Eds., Groundwater ecology. Academic Press, San Diego, CA, pp. 477 – 504.
Oberlin, G. E., Blinn, D. W. 1997. The effect of temperature on the
metabolism and behavior of an endemic amphipod, Hyalella
montezuma from Montezuma Well, Arizona, USA. Freshwater
Biology 37:55 – 59.
Ogarkov, O. B., Kamaltynov, R. M., Belikov, S. I., Shcherbakov, D. Y
1997. Phylogenetic relatedness of the Baikal Lake endemial amphipodes (Crustacea, Amphipoda) deduced from partial nucleotide sequences of the cyctochrome oxidase subunit III genes.
Molecular Biology 31:24 – 29.
Okland, K. A., Okland, J. 1985. Factor interaction influencing the
distribution of the fresh-water shrimp Gammarus. Oecologia
66:364 – 367.
Olyslager, N. J., Williams, D. D. 1993. Microhabitat selection by the
lotic amphipod Gammarus pseudolimneaus Bousfield — mechanisms for evaluating local substrate and current suitability.
Canadian Journal of Zoology 71:2401 – 2409.
Palmer, M., Covich, A. P., Finlay, B. J., Gibert, J., Hyde, K. D., Johnson, R. K., Kairesalo, T., Lake, S., Lovell, C. R., Naiman, R. J.,
Ricci, C., Sabater, F., Strayer, D. 1997. Biodiversity and ecosystem processes in freshwater sediments. Ambio 26:571 – 577.
Panov, V. E., McQueen, D. J. 1998. Effects of temperature on individual growth rate and body size of a freshwater amphipod.
Canadian Journal of Zoology 76:1107 – 1116.
Parker, J. I. 1980. Predation by Mysis relicta on Pontoporeia hoyi: a
food chain link of potential importance in the Great Lakes.
Journal of Great Lakes Research 6:164 – 166.
Peck, S. B. 1998. A summary of diversity and distribution of the obligate cave-inhabiting fauna of the United States and Canada.
Journal of Cave and Karst Studies 60:18 – 26.
Peckarsky, B. L., Frassinet, P. R., Penton, M. A., Conklin, D. J., Jr.
1990. Freshwater macroinvertebrates of northeastern North
America. Cornell Univ. Press, Ithaca, New York. 422 pp.
Pennak, R. W. 1989. Freshwater invertebrates of the United States:
protozoa to mollusca, 3rd ed., Wiley, New York. 628 p.
Pixell Goodrich, H. P. 1934. Reactions of Gammarus to injury and
disease, with notes on microsporidial and fungoid disease.
Quarterly Journal of the Microscopy Society 72:325 – 353.
Ricciardi, A., Rasmussen, J. B. 1999. Extinction rates of North American freshwater faunas. Conservation Biology 13:1220 – 1222.
Richardson, H. 1905. A monograph of the isopods of North America. Bulletin of the U.S. National Museum 54:1 – 727.
Roberts, G. N., Chubb, J. C. 1998. The distribution and location of
the symbiont Lagenophyrs aselli on the freshwater isopod Asellus aquaticus. Freshwater Biology 39:671 – 677.
Roman Contreras, R. 1996. A new species of Probopyrus (Isopoda,
Bopyridae), parasite of Macrobrachium americanum Bate, 1868
(Decapoda, Palaemonidae). Crustaceana 69:204 – 210.
Rudstam, L. G., Melnik, N. G., Timoshkin, O. A., Hansson, S.,
Pushkin, S. V., Nemov, V. 1992. Diel dynamics of an aggregation of Macrohectopus branickii (Dyb) (Amphipoda, Gammaridae) in the Barguzin Bay, Lake Baikal, Russia. Journal of Great
Lakes Research 18:286 – 297.
Rudstam, L. G., Hetherington, A. L., Mohammadian, A. M. 1999.
Effect of temperature on feeding and survival of Mysis relicta.
Journal of Great Lakes Research 25:363 – 371.
Ruff, H., Maier, G. 2000. Calcium carbonate deposits reduce predation on Gammarus fossarum from salamander larvae. Freshwater Biology 43:99 – 105.
Sarbu, S., Kane, T. C., Culver, D. C. 1993. Genetic structure and morphological differentiation: Gammarus minus (Amphipoda, Gammaridae) in Virginia. American Midland Naturalist 129:145 – 152.
Sarvala, J. 1986. Interannual variation of growth and recruitment
in Pontoporeia affinis (Lindstrom) (Crustacea, Amphipoda) in
relation to abundance fluctuations. Journal of Experimental
Marine Ecology 101:41 – 59.
19. Introduction to the Subphylum Crustacea
Sastry, A. N. 1983. Ecological aspects of reproduction, in: Vernberg,
F. J., Vernberg, W. B., Eds., The biology of Crustacea, Vol. 8.
Academic Press, New York, pp. 179 – 270.
Schmitt, W. L. 1965. Crustaceans, University of Michigan Press, Ann
Arbor, 204 p.
Schmitz, E. H. 1992. Amphipoda, in: Harrison, F. W., Ed., Microscopic anatomy of invertebrates. Vol. 9 Crustacea. Wiley – Liss,
New York, pp. 443 – 528.
Schmitz, E. H., Scherrey, P. M. 1983. Digestive anatomy of Hyalella
azteca (Crustacea, Amphipoda). Journal of Morphology
175:91 – 100.
Schram, F. R. 1982. The fossil record and evolution of Crustacea, in:
Abele, L. G., Ed., The biology of Crustacea, Vol. 1. Academic
Press. New York, pp. 93 – 174.
Schram, F. R. 1986. Crustacea, Oxford University Press, New York,
606 p.
Schram, F. R., Yager, J., Emerson, M. J. 1986. Remipedia. Part I. Systematics. Memoirs of the San Diego Society of Natural History
15:1 – 60.
Schwartz, S. S. 1992. Benthic predators and zooplankton prey: predation by Crangonyx shoemakeri (Crustacea: Amphipoda) on
Daphnia obtusa (Crustacea: Cladocera). Hydrobiologia 237:
25 – 30.
Seale, D. B., Binowski, F. P. 1988. Vulnerability of early life intervals
of Coregonus hoyi to predation by a freshwater mysid. Mysis
relicta. Environmental Biology of Fishes 21:117 – 126.
Sergestrale, S. G. 1970. Light control of the reproductive cycle of
Pontoporcia affinis Lindstrom (Crustacea: Amphipoda). Journal
of Experimental Marine Biology and Ecology 5:272 – 275.
Shafir, A., Oldewage, W. H. 1992. Dynamics of a fish ectoparasite
population- opportunistic parasitism in Argulus japonicus
(Branchiura). Crustaceana 62:50 – 64.
Shea, M. A., Makarewicz, J. C. 1989. Production, biomass, and
trophic interactions of Mysis relicta in Lake Ontario. Journal of
Great Lakes Research 15:223 – 232.
Sherbakov, D., R. M., Ogarkov, O. B., Verheyen, E. 1998. Patterns
of evolutionary change in Baikalian gammarids inferred from
DNA sequences (Crustacea, Amphipoda). Molecular Phylogenetics and Evolution 10:160 – 167.
Sherman, R. K., Lasenby, D. C., Hollet, L. 1987. Influence of oxygen
concentration on the distribution of Mysis relicta Loven in a eutrophic temperate lake. Canadian Journal of Zoology 65:
2646 – 2650.
Short, T. M., Holomuzki, J. R. 1992. Indirect effects of fish on foraging behavior and leaf processing by the isopod Lirceus fontinalis. Freshwater Biology 27:91 – 97.
Shuster, S. M. 1981. Life history characteristics of Thermosphaeroma
thermophi!um, the Socorro isopod (Crustacea: Peracarida). Biological Bulletin 161:291 – 302.
Sieg, J. 1983. Evolution of the Tanaidacea, in: Schram, F. R., Ed.,
Crustacean phylogeny. Balkema, Rotterdam, pp. 229 – 256.
Siegfried, C. A. 1985. Life history, population dynamics and production of Pontoporeia hoyi (Crustacea, Amphipoda) in relation to
the trophic gradient of Lake George, New York. Hydrobiologia
122:175 – 180.
Siegismund, H. R., Muller, J. 1991. Genetic structure of Gammarus
fossarum populations. Heredity 66:419 – 436.
Simon, K. S., Buikema, A. L. 1997. Effects of organic pollution on an
Appalachian cave: changes in macroinvertebrate populations.
American Midland Naturalist 138:387 – 401.
Sket, B. 1996. The ecology of anchihaline caves. Trends in Ecology
and Evolution 11:221 – 225.
Sket, B. 1999a. High biodiversity in hypogean waters and its endangerment — the situation in Slovenia, the Dinaric karst, and Europe. Crustaceana 72:767 – 779.
807
Sket, B. 1999b. The nature of biodiversity in hypogean waters
and how it is endangered. Biodiversity and Conservation 8:
1319 – 1338.
Sly, P. G., Christie, W. J. 1992. Factors influencing densities and distributions of Pontoporeia hoyi in Lake Ontario. Hydrobiologia
235:321 – 352.
Smock, L. A., Harlowe, K. L. 1983. Utilization and processing of
freshwater wetland macrophytes by the detritivore Asellus
forbesi. Ecology 64:1556 – 1565.
Smock, L. A., Stoneburner, D. L. 1980. The response of macroinvertebrates to aquatic macrophyte decomposition. Oikos 33:
397 – 403.
Song, K. H., Breslin, V. T. 1998. Accumulation of contaminant metals in the amphipod Diporeia spp. in western Lake Ontario.
Journal of Great Lakes Research 24:949 – 961.
Song, K. H., Breslin, V. T. 1999. Accumulation and transport of sediment metals by the vertically migrating opossum shrimp, Mysis
relicta. Journal of Great Lakes Research 25:429 – 442.
Sparkes, T. C. 1996a.The effects of size-dependent predation risk on
the interaction between behavioral and life history traits in a
stream-dwelling isopod. Behavioral Ecology and Sociobiology
39:411 – 417.
Sparkes, T. C. 1996b. Effects of predation risk on population variation in adult size in a stream dwelling isopod. Oecologia
106:85 – 92.
Spencer, C. N., McClelland, B. R., Stanford, J. A. 1991. Shrimp introduction, salmon collapse, and eagle displacement: cascading
interactions in the food web of a large aquatic ecosystem. BioScience 41:14 – 21.
Spencer, C. N., Potter, D. S., Bukantis, R. T., Stanford, J. A. 1999.
Impact of predation by Mysis relicta on zooplankton in Flathead Lake, Montana, USA. Journal of Plankton Research
21:51 – 64.
Stanford, J. A., Ward, J. V. 1988. The hyporheic habitat of river
ecosystems. Nature 335:64 – 66.
Strayer, D. 1999. Invasion of fresh waters by saltwater animals.
Trends in Ecology and Evolution 14: 448 – 449.
Strayer, D. L., May, S. E., Nielsen, P., Wollheim, W., Hausam, S.
1997. Oxygen, organic matter, and sediment granulometry as
controls on hyporheic animal communities. Archiv fur Hydrobiologie 140:131 – 144.
Stephens, D. W. 1990. Changes in lake levels, salinity and the biological community of Great Salt Lake (Utah, USA), 1847 – 1987.
Hydrobiologia 197:139 – 146.
Stewart, B. A. 1992. Morphological and genetic differentiation between allopatric populations of a freshwater amphipod. Journal
of Zoology (London) 228:287 – 305.
Stewart, B. A. 1993. The use of protein electrophoresis for determining species boudaries in amphipods. Crustaceana 65:265 – 277.
Stock, F. 1995. The ecological and historical determinants of crustacean diversity in groundwaters, or : why are these so many
species? Memoirs of Biospeology 22:139 – 160.
Stock, J. H., Iliffe, T. M. 1990. Amphipod crustaceans from anchialine cave waters of the Galapagos Islands. Zoological Journal of
the Linnean Society 98:141 – 160.
Stock, J. H., Longley, G. 1981. The generic status and distribution of
Monodella rexana, Maguire, the only known North American
thermosbaenacean. Proceedings of the Biological Society of
Washington 94:569 – 578.
Strong, D. R. 1972. Life history variation among populations of an
amphipod (HyaleIla azteca). Ecology 53:1103 – 1111.
Strong, D. R., Jr. 1973. Amphipod amplexus, the significance of ecotypic variation. Ecology 54:1383 – 1388.
Sutcliffe, D. W. 1971. Regulation of water and some ions in gammarids (Amphipoda). I. Gammarus duebeni Lilljeborg from
808
A. P. Covich and J. H. Thorp
brackish water and fresh water. Journal of Experimental Biology 55:325 – 344.
Sutcliffe, D. W. 1974. Sodium regulation and adaptation to fresh
water in the isopod genus Asellus. Journal of Experimental
Biology 61:719 – 736.
Sutcliffe, D. W., Carrick, T. R., Willoughby, L. G. 1981. Effects of
diet. body size, age and temperature on growth rates in the amphipod Gammarus pulex. Freshwater Biology 11:183 – 214.
Tadini, G. V., Tadini, E. A., Colangelo, M. 1988. The life history of
Asellus aquaticus (L.) explains its geographical distribution.
Verhandlungen Internationale Vereinigung fur Theoretische und
Angewandte Limnologie 23:2099 – 2106.
Taylor, C. A. 2000. Preserving North America’s unique crayfish
fauna, in: Abell, R. A., Olson, D. M., Dinerstein, E., Hurley,
P. T., Diggs, J. T., Eichbaum, W., Walters, S., Wettengel, W.,
Allnutt, T., Loucks, C. J., Hedau, P., Eds., Freshwater ecoregions of North America. A conservation assessment. Island
Press, Washington, D C, pp. 36 – 37.
Thomas, P. E., Blinn, D. W., Keim, P. 1997. Genetic and behavioural
divergence among desert spring amphipod populations. Freshwater Biology 38:137 – 143.
Thomas, P. E., Blinn, D. W., Keim, P. 1998. Do xeric landscapes increase genetic divergence in aquatic ecosystems? Freshwater Biology 40:587 – 593.
Threlkeld, S. T., Chiavelli, D. A., Willey, R. L. 1993. The organization of zooplankton epibiont communities. Trends in Ecology
and Evolution 8:317 – 321.
Thybaud, E., LeBrac, S. 1988. Absorption and elimination by Asellus
aquaticus (Crustacea, Isopoda). Bulletin of Environmental Contamination and Toxicity 40:731 – 735.
Toman, M. J., Dall, P. C. 1998. Respiratory levels and adaptations in
four freshwater species Gammarus (Crustacea: Amphipoda). International Review of Hydrobiology 83:251 – 263.
Vainola, R. 1990. Molecular time scales for evolution in Mysis and
Pontoporeia. Annales Zoologici Fennici 27:211 – 214.
Vainola, R., Kamaltynov, R. M. 1999. Species diversity and speciation in the endemic amphipods of Lake Baikal: molecular evidence. Crustaceana 72:945 – 956.
Vainola, R., Varvio, S. L. 1989. Molecular divergence and evolutionary relationships in Pontoporeia (Crustacea: Amphipoda).
Canadian Journal of Fisheries and Aquatic Sciences 46:
1705 – 1713.
Vainio, J., Jazdzewski, K., Vainola, R. 1995. Biochemical systematic
relationships among the freshwater amphipods Gammarus
varsoviensis, Gammarus lacustris and Gammarus pulex. Crustaceana 68:687 – 694.
Vernberg, W. B., Vernberg, F. J. 1983. Freshwater adaptations, in:
Vernberg, F. J., Vernberg, W. B., Eds., The biology of Crustacea,
Vol 8. Academic Press, New York, pp. 335 – 363.
Wagner, V. T., Blinn, D. W. 1987. A comparative study of the maxillary setae for two coexisting species of Hyalella (Amphipoda), a
filter feeder and a detritus feeder. Archiv fur Hydrobiologia
109:409 – 419.
Waters, T. F., Hokenstrom, J. C. 1980. Annual production and drift
of the stream amphipod Gammarus pseudolimnaeus in Valley
Creek, Minnesota. Limnology and Oceanography 25:700 – 710.
Watling, L. 1981. An alternative phylogeny of peracarid crustaceans.
Journal of Crustacean Biology 1:201 – 210.
Wellborn, G. A. 1994. Size-biased predation and prey life histories: a
comparative study of freshwater amphipod populations. Ecology 75:2104 – 2117.
Wellborn, G. A. 1995. Determinants of reproductive success in freshwater amphipod species that experience different mortality
regimes. Animal Behaviour 50:353 – 363.
Wellborn, G. A. 2000. Selection on a sexually dimorphic trait in ecotypes within the Hyalella azteca complex (Amphipoda: Hyalellidae). American Midland Naturalist 143:212 – 225.
Wells, L. 1960. Seasonal abundance and vertical movements of
planktonic Crustacea in Lake Michigan. U.S. Fish and Wildlife
Service Fishery Bulletin 60:343 – 369.
Wen, Y. H. 1993. Sexual dimorphism and mate choice in Hyalella
azteca (Amphipoda). American Midland Naturalist 129:
153 – 160.
Whitehurst, I. T. 1991.The Gammarus – Asellus ratio as an index of
organic pollution. Water Research 25:333 – 339.
Wickstrom, C. E., Castenholz, R. W. 1973. Thermophilic ostracod:
aquatic metazoan with the highest known temperature tolerance. Science 181:1063 – 1064.
Wilhelm, F. M. 1996. Predation on Mysis relicta by slimy sculpins
(Cottus cognatus) in southern Lake Ontario — comment. Journal of Great Lakes Research 22:119 – 120.
Wilhelm, F. M., Schindler, D. W. 2000. Reproductive strategies of
Gammarus lacustris (Crustacea: Amphipoda) along on elevational gradient. Functional Ecology 14: 413 – 422.
Williams, D. D. 1990. A field study of the effects of water temperature, discharge and trout odor on the drift of stream invertebrates. Archiv fur Hydrobiologie 119:167 – 181.
Williams, D. D., Moore, K. A. 1985. The role of semiochemicals
to benthic community relationships of the lotic amphipod
Gammarus pseudolimnaeus: a laboratory analysis. Oikos 44:
280 – 286.
Williams, W. D. 1970. A revision of North American epigean species
of Asellus (Crustacea: Isopoda). Smithsonian Contributions to
Zoology 49:1 – 80.
Williams, W. D. 1998. Salinity as a determinant of the structure
of biological communities in salt lakes. Hydrobiologia 381:
191 – 201.
Willoughby, L. G., Sutcliffe, D. W. 1976. Experiments on feeding
and growth of the amphipod Gammarus pulex (L.) related to
its distribution in the River Duddon. Freshwater Biology
6:577 – 586.
Wilson, C. B. 1944. Parasitic copepods of the United States National
Museum. Proceedings of the U.S. National Museum 94:529 – 582.
Winnell, M. H., White, D. S. 1984. Ecology of shallow and deep water populations of Pontoporeia hoyi (Smith) (Amphipoda) in
Lake Michigan. Freshwater Invertebrate Biology 3:118 – 138.
Wisenden, B. D., Cline, A., Sparkes, T. C. 1999. Survival benefit to
antipredator behavior in the amphipod Gammarus minus
(Crustacea: Amphipoda) in response to injury-released chemical cues from conspecifics and heterospecifics. Ethology 105:
407 – 414.
Wollheim, W. M., Lovvorn, J. R. 1995. Salinity effects on macroinvertebrate assemblages and waterbird food webs in shallow lakes of
the Wyoming high plains. Hydrobiologia 310:207 – 223.
Wollheim, W. M., Lovvorn, J. R. 1996. Effects of macrophyte
growth forms on invertebrate communities in saline lakes of
Wyoming high plains. Hydrobiologia 323:83 – 96.
Wooster, D. E. 1998. Amphipod (Gammarus minus) responses to
predators and predator impact on amphipod density. Oecologia
115:253 – 259.
Wudkevich, K., Wisenden, B. D., Chivers, D. P., Smith, R. J. F. 1997.
Reactions of Gammarus lacustris to chemical stimuli form natural predators and injured conspecifics. Journal of Chemical
Ecology 23:1163 – 1173.
Wurtsbaugh, W. A., Berry, T. S. 1990. Cascading effects of decreased
salinity on the plankton, chemistry, and physics of the Great
Salt Lake (Utah). Canadian Journal of Fisheries and Aquatic
Sciences 47:100 – 109.
19. Introduction to the Subphylum Crustacea
Yager, J. 1981. Remipedia. a new class of Crustacea from a marine cave
in the Bahamas. Journal of Crustacean Biology 1: 328 – 333.
Yager, J. 1987. Cryptocorynectes haptodiscus and Speleonectes benjamini from anchialine caves in the Bahamas, with remarks on
distribution and ecology. Proceedings of the Biological Society
of Washington 100:302 – 320.
Yamaguti, S. 1963. Parasitic Copepoda and Branchiura of fishes. Wiley Interscience, New York. 1104 pp.
Yampolsky, L. Y., Kamaltynov, R. M., Ebert, D., Filatov, D. A.,
Chernyck, V. I. 1994. Variation of allozyme loci in endemic
809
gammarids in Lake Baikal. Biological Journal of the Linnean
Society 53:309 – 323.
Yoshii, K., Melnik, N. G., Timoshkin, O. A., Bondarenko, N. A.,
Anoshko, P. N., Yoshioka, T., Wada, E. 1999. Stable isotope
analyses of the pelagic food web in Lake Baikal. Limnology and
Oceanography 44:502 – 511.
Zimmer-Faust, R. K. 1989. The relationship between chemoreception
and foraging behavior in crustaceans. Limnology and Oceanography 34:1367 – 1374.
This Page Intentionally Left Blank
20
OSTRACODA
L. Denis Delorme1
National Water Research Institute
Canada Centre for Inland Waters
Burlington, Ontario, L7R 4A6
I. Introduction
II. Anatomy and Physiology
A. External Shell Morphology
B. Internal Shell Morphology
C. Body and Appendage
Morphology
D. Internal Anatomy and Physiology
III. Life History
A. Egg
B. Reproduction
C. Embryology
D. Hatching and Seasonality
E. Molting
F. Competition, Coexistence, and
Survival
IV. Distribution
A. Benthic Swimmers
B. Benthic Nonswimmers
V. Chemical Habitats
A. Chemical Water Types
B. Salinity and Solute Composition
C. Hydrogen Ion Concentration
D. Dissolved Oxygen
E. Carbonate
VI. Physical Habitat
A. Temperature
B. Lakes, Ponds, and Streams
C. Bromeliads
D. Cave and Subterranean Habitats
I. INTRODUCTION
The class Ostracoda in the Crustacea contains both
marine and freshwater forms. The subclass Podocopa
has one order, Podocopida in which two of the suborders (Metacopina, Podocopina) contain all of the fresh1
Current address: 621 Auburn Crescent, Burlington, Ontario,
Canada L7L 5B3
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
VII.
VIII.
IX.
X.
XI.
XII.
XIII.
XIV.
E. Sediment – Water Interface,
Particle Size, Food
Physiological and Morphological
Adaptations
A. Eggs
B. Resting Stage and Torpidity
C. Coloration
D. Appendages
Behavioral Ecology
Foraging Relationships
Population Regulation
A. Suitability of Habitat
B. Toxicity of Herbicides and Pesticides
C. Ultraviolet-B Radiation
D. Parasitism
Collecting and Rearing Techniques
A. Collecting and Preservation
B. Rearing Techniques
Uses of Freshwater Ostracoda
Current and Future Research Problems
Classification of Ostracodes
A. Taxonomic Controversies
B. Classification of the Order
Podocopida
C. Taxonomic Key to Genera of the
Superfamily Cypridoidea
D. Taxonomic Key to Genera of the
Superfamily Cytheroidea
Literature Cited
water ostracodes (Bowman and Abele, 1982; Schram,
1986).
Ostracodes are the oldest known microfauna. Their
preservation as fossils is attributed to the calcitic shell
present in most species. The first fossil representatives
are reported from Cambrian marine sediments (Moore,
1961; Maddocks, 1982). The earliest freshwater ostracode species came from coal-forming swamps, ponds,
and streams of early Pennsylvanian age (Benson, 1961).
811
812
L. D. Delorme
According to Moore (1961), the Podocopina evolved
from Podocopida stock during the Ordovician period,
and then branched out into the Cytheroidea and the
Cypridoidea during the Devonian period (Maddocks,
1982). The Darwinulidae evolved during the Carboniferous period (Maddocks, 1982).
Freshwater ostracodes are free living except the
Entocytheridae (Hoff, 1942) which are commensal
upon crayfish. This chapter will concentrate on freshwater ostracode genera found in North America. To
date there have been 56 genera and about 420 species
identified for this continent.
Most references older than 1900 have been deleted
from the second edition. Those interested in the older
references are referred to the original edition.
II. ANATOMY AND PHYSIOLOGY
A. External Shell Morphology
The visceral mass is completely encased within a
chitinous and a low-magnesium calcite exoskeleton
(carapace). The only exception is the entocytherid
group of ostracodes which lacks calcium in the exoskeleton. The exterior surface of the exoskeleton (cuticle) is made up of chitin (Fig. 1) (Harding, 1964). The
outer chitinous coating of the epidermis is the only covering over the calcitic shell. The soft epidermal tissue
lining the interior part of the carapace is composed of
the inner and outer epidermis. The epidermal cells of
the outer lamella (Fig. 1) are larger and more irregular
in shape than the inner epidermal cells. Subdermal cells
occur between the inner and outer epidermal cells as do
the liver, ovaries, and testes. The epidermis is contained
within a thin chitinous lining. When the ostracode
molts, the complete exoskeleton is shed leaving only
the visceral mass and the soft tissue or epidermis. The
epidermis is continuous with the visceral mass in the
dorsal region of the body.
When the ostracode molts, the space between the
epidermal cells of the outer and inner layer is empty.
The shells are secreted by the cells of the outer lamellae
(Turpen and Angell, 1971). The immediate source of
calcium and magnesium carbonate is thought to be
stored in the body and not obtained directly from the
water (Fassbinder, 1912). Fassbinder has also shown
that ostracodes fed with crushed calcium carbonate
from snail shells developed thicker carapaces with more
pronounced marginal ornamentation. He concluded
that food is an important source of carbonate. Turpen
and Angell (1971), however, found that calcium dissolved in water is the source of calcium for the shell.
They used dissolved 45Ca as a tracer (pH 7.4, 23°C) to
FIGURE 1
Diagrammatic cross section of the shell and the
enclosing dermal layers, vestibule located between the outer lamella
and the shell portion of the inner lamella in the anterior portion of
the shell. [From Kesling, 1951a, b. Figure redrawn by J. L. Delorme.]
show that calcium is not resorbed from the shells prior
to molting and that ostracodes do not store calcium in
their bodies. Chivas et al., (1985) ran laboratory experiments on the Australian Mytilocypris henricae and
found that temperature influenced the rate of shell calcification. Roca and Wansard (1997) also found the rate
of development to be linked to temperature. Using Herpetocypris brevicaudata they were able to show that below 15°C the Ca content of valves is strongly diminished and the length of time for calcification was longer
(17 days). The optimum temperature was 23°C with the
fastest rate of calcification (3 – 7 days). Bodergat (1978)
demonstrated that phosphorus was an integral part of
freshwater ostracode shells in all specimens investigated
with a microprobe. She further noted a significant positive correlation between the amount of shell phosphorus and the habitat temperature of the ostracode.
The calcareous shell is the most obvious structure
when the organism is viewed under a microscope. The
shell is divided into two parts, the outer lamella and
the duplicature (Fig. 1). The outer lamella is the major
part of the shell; whereas the duplicature is the calcified
part of the inner lamella bordering the posterior, ventral, and anterior margin. It forms a part of the free
margin and projects toward the center of the carapace.
The outer lamella may contain pores through which
20. Ostracoda
project setae (Fig. 2), that are sensitive to touch. The
duplicature is welded to the outer lamella (Fig. 3)
around the margins, by a strip of chitin, which joins
with the chitin coating and the chitin lining. This juncture is referred to as the line of concrescence or the distal line of adhesive strip (Fig. 1). Within the chitinous
strip, radial or marginal pore canals are found (Fig. 2).
These are very fine tubes through which setae pass. The
inner free margin of the duplicature is referred to as the
inner margin (Fig. 1). When the duplicature extends beyond the line of concrescence toward the center of the
shell, a shelf is formed (Fig. 3). The space between
the shelf and the outer lamella is referred to as the
vestibule (Fig. 1). Several structures are found on the
duplicature which are very important in taxonomy
down to the species level (Sylvester-Bradley, 1941).
Pustules, teeth, or crenulations (Fig. 3) may appear
on the outer margin of the duplicature. A combination
of septa, lists, and grooves are found on the duplicature. Some of these are shown in Fig. 1 (Kesling,
1951a, b). The shell may contain additional structures
on the outer lamella such as lateral depressions called
sulci (Fig. 30). Raised areas variously termed alae,
knobs (Fig. 30), or pustules (Fig. 4), depending on their
shape, size, and orientation, may also be found on the
shell surface. The surface of the shell may also be pitted
(Fig. 5), punctate, wrinkled (Fig. 6), or have a reticulate
surface (Fig. 7). These ancillary structures of the shell
are very often used for generic and specific identification (see Sylvester-Bradley and Benson, 1971, for terminology). Some authors, notably Benson (1974, 1981),
have discussed the roles of ornamentation, form, function, and architecture of the ostracode shell.
Intraspecific morphological diversity of the shell
may be pronounced. Limnocythere itasca has short recurved alae on its lateral surfaces. The individuals identified from Canada (Delorme, 1971a) and northern
Minnesota (Forester et al., 1987) appear to be characteristic of this species. R. M. Forester (personal communication) identified L. itasca from the Laguna de
Texcoco, Mexico, which have very long and delicate
recurved spines or alae. Intermediate forms have yet to
be recovered. In another example, the caudal process of
the posteroventral portion of the right valve of the female Candona caudata (may belong to Fabaeformiscandona; see Meisch, 1996, not confirmed for North
America) may be blunt (Delorme, 1978) or sharp ( C.
novacaudata, Benson and MacDonald, 1963). Both
forms and intermediate morphs are found in Lake Erie.
B. Internal Shell Morphology
Structures on the internal surface of the shell may
also be used in ostracode systematics. Many ostracode
813
muscles are attached directly to the calcareous shells.
The chitinous exoskeleton does not provide sufficient
rigidity to anchor many of the muscles. The points of
attachment of these muscles leave scars as raised or depressed areas on the interior of the shell (Figs. 8-11).
The closing or adductor muscles form a distinctive pattern of scars as do the mandibular muscles (Benson,
1967; Benson and MacDonald, 1963; Smith, 1965).
Traces of the ovaries and testes can also be seen in the
posterior of the shell (see Section II.D).
C. Body and Appendage Morphology
Ostracodes have their body (visceral mass and appendages) suspended from the dorsal region in an elongate, chitinous pouch. The ostracode has a shortened
body with a slight constriction in the midregion that
separates the head from the thorax (Kesling, 1951a).
There is no abdomen. Instead, the posterior of the
body tapers off bluntly and ends in a pair of preanal
furcae. The head region contains four pairs of appendages which are used for swimming, walking, and
feeding. The thoracic region features two pairs (three
pairs according to some authors) of appendages used
or adapted for feeding, creeping, and cleaning of the
shells (Broodbakker and Danielopol, 1982). There is
some debate as to whether the maxilla are cephalic or
thoracic appendage (Meisch, 1996) Ancillary cuticular,
structures such as setae, claws, and pseudochaetae,
found on most limbs, are recognized as important
in functional morphology and systematics (Broodbakker and Danielopol, 1982). Appendages are shown
in Figs. 9 – 12.
The head region is composed of the forehead, the
upper lip, and the hypostome. The four pairs of appendages in the head region are the first antennae
(antennules), second antennae (antennae), mandibles,
and maxillae. When the distal four podomeres of the
first antennae have very long, plumose setae, the ostracode is a good swimmer (e.g., Cypridopsis vidua). The
first antennae are curved up and backward. If these setae are absent, as in the darwinulids (Fig. 10) and the
limnocytherids (Fig. 11), the ostracode cannot swim.
The second antennae extend down and curve backward. These appendages are robustly constructed and
have a strong claw for walking and climbing. Swimming setae may be present on the first podomere of the
endopodite. These structures are used for rowing when
the ostracode is swimming. In most species, welldeveloped mandibles occur between the upper lip and
the hypostome. The dorsal tips of the mandibular palps
are attached to the interior of the shell by muscles. The
ventral portion of the palps terminate in heavily
toothed mandibles; these teeth grind the food before it
FIGURE 2 Shell surface of Cyclocypris ampla Furtos showing normal pore canals with funnels (without seta)
and without funnels (with seta), and dimples. FIGURE 3 Anterior margin of Cyprinotus glaucus Furtos
showing crenulations, duplicature with position of radial pore canals, and calcified inner lamella forming a
shelf. FIGURE 4 Anterior margin of Cyprinotus glaucus Furtos with pustules on the right valve, left valve
beneath. FIGURE 5 Reticulate exterior structure on Limnocythere itasca Cole. FIGURE 6 Anastomosing
structure (lines) on the surface of Cypria turneri, note normal pore canals. FIGURE 7 Reticulate structure
with superimposed mamilliary pustules on Paracandona euplectella Brady and Norman. FIGURE 8
Distinctive muscle scar pattern of Darwinula.
814
20. Ostracoda
815
FIGURE 10 Sketch of the internal morphology of a female
Darwinula stevensoni (Brady and Robertson) (Darwinulidae) from
Kesling. First thoracic leg is referred to as the maxilla or maxillar
endopodite and the second and third thoracic legs as the first and
second thoracic legs in the text. [Figure redrawn by J. L. Delorme.]
FIGURE 9 Sketch of the internal morphology of (a) male (b) female
Candona suburbana Hoff (Cyprididae) from Kesling. First thoracic
leg is referred to as the maxilla or maxillar endopodite and the
second and third thoracic legs as the first and second thoracic legs in
the text. [Figure redrawn by J. L. Delorme.]
FIGURE 11 Sketch of the internal morphology of a male Limnocythere sanctipatricii (Brady and Robertson) (Cytheridae) from
Kesling. First thoracic leg is referred to as the maxilla or maxillar
endopodite and the second and third thoracic legs as the first and
second thoracic legs in the text. [Figure redrawn by J. L. Delorme.]
is ingested. A respiratory (branchial) plate extends
from the mandibular palp. On the ventral portion of
the head is located a keel-shaped structure called the
hypostome which forms the mouth. Rake-shaped organs are situated at the rear of the mouth. They consist
of chitin shafts with terminal toothed structures and
are used for straining and feeding. The fourth and final
set of cephalic appendages are the maxillae. The maxillae, located on either side of the hypostome posterior
of the mandibles, are made up of several cylindrical
masticatory processes (maxillula). The maxillae pass
the small particles toward the mouth. What were previously referred to as the first thoracic legs of most
cyprids are now referred to as part of the maxillae. In
other groups, this part of the maxillae (formerly the
FIGURE 12
Generalized sketch of the internal morphology of a
male entocytherid (Cytheridae) from Hart and Hart. First thoracic
leg is referred to as the maxilla or maxillar endopodite and the
second and third thoracic legs as the first and second thoracic legs in
the text. [Figure redrawn by J. L. Delorme.]
816
L. D. Delorme
first thoracic legs) are modified in sexual ostracodes
into prehensile palps (Fig. 9a) used in grasping the female shell during copulation. There are two pairs of
thoracic appendages. The first thoracic legs (previously
called the second thoracic legs) are robustly developed
and used for walking, climbing, and clinging onto surfaces. The second thoracic leg (formerly called the third
thoracic legs), or cleaning legs, are the most flexible
and well-muscled appendages. The end of these legs, in
most ostracodes, are modified pinchers used to grasp
and remove foreign objects from between the shells. In
the entocytherid group, the modified maxillae and the
two pairs of thoracic legs (Fig. 12) are modified terminally as hooks to hold onto the gills of the host crayfish
(Hart and Hart, 1974).
Morphological diversity can be seen in the maxillae of ostracodes. Most notable are the first thoracic
legs. These can be modified for feeding, for use during
copulation (prehensile palps), or for walking. The commensal entocytherids have modified the maxillae and
two thoracic legs for grasping onto the gills of the host.
Paired hemipenes are found in syngamic species behind the third thoracic legs on the ventral side of the
thorax. Paired uterine openings lie behind and inside
the vaginal openings of sexual and asexual females.
The distal part of the thorax terminates in paired furcae. In some species, these are whip-shaped structures
while in others they are well developed and armed at
the end with long, serrated claws. Rome (1969) and
Triebel (1953) consider the furcal attachment to be an
important biocharacter to be used in systematics.
Sexual dimorphism is common in ostracodes. Determination of gender in most cases can be determined
visually with a microscope. This is done by observing
the presence / absence of testes and traces of the testes
on the inner surface of the shell, Zenker’s organs (see
Section II.D), modification of the male maxillae (formerly called the first thoracic leg) into prehensile palps,
hemipenes, and ovaries. For many species, it is necessary to separate the valves to view these organs. Except
in cyprinotids, the male is usually larger than the female. In candonids, only the male has an anteroventral
notch on the shell.
D. Internal Anatomy and Physiology
The circulatory system of freshwater ostracodes
lacks both heart and gills. Gaseous exchange is through
the entire surface of the body and particularly the
membranous inner lamella of the exoskeleton. The respiratory plates of some appendages move and renew
oxygenated water past the inner surfaces of the organisms. Large respiratory cells are known to occur in the
inner lamella, forming a respiratory epithelium over the
valve cavity which forms a blood sinus (Bernecker,
1909).
The digestive system of ostracodes consists of the
mouth, esophagus, stomach, intestine, rear gut, and anus.
The mouth is a large opening with the large teeth of the
mandibular palps on either side. A gland, probably a salivary gland, opens into the mouth. The esophagus is very
muscular and leads into the distended stomach. The hepatopancreas (liver) is found in the epidermal layers next
to the shells and empties into the anterior part of the
stomach. Most of the digestion takes place in the stomach where the food is formed into balls, passes from the
stomach through the rear gut for absorption of the nutrients, and exits the anus as fecal pellets.
Approximately eight excretory glands, including
the liver, have been identified in freshwater ostracodes.
McGregor (1967) showed that for Chlamydotheca arcuata some food particles pass through the hepatopancreas, aiding in the digestion of food. Most glands have
been studied in considerable detail (Bergold, 1910;
Fassbinder, 1912; Schreiber, 1922; Cannon, 1925;
Rome, 1947; Kesling, 1951a, 1965).
The nervous system of the ostracode is composed
of a cerebrum (protocerebrum, deutocerebrum, and tritocerebrum; Turner, 1896; Hanström, 1924; Weygoldt,
1961), circumesophageal ganglion, and a ventral chain
of fused ganglia (Kesling, 1951a). The first and second
antennae, mandibular palps, and maxillae have small
ganglia which connect to the central nervous system.
The ostracode has a single median eye made up of
three optic cups each containing lenses. The number of
cells composing a lens is variable (Novikoff, 1908;
Rome, 1947). The black opaque eye lies in the anterior
part of the body just below the rim of the shells. When
the shells are open, the animal can detect shapes and
motion; however, when the shells are closed it can only
distinguish light intensity. Subterranean-interstitial (hypogean) ostracodes are blind (Danielopol, 1980a).
The ovaries are found in the epidermis next to the
shell as a continuous sequence of gametes. The ova are
fairly large in the last instar of many species, forcing the
chitin coating of the epidermis into the space normally
occupied by the shell. When the new shell is secreted, a
trace of the ovaries is commonly seen as part of the
shell’s inner structure in the Candoninae. The testes are
similarly placed and appear as traces on the inner and
posterior region of the shell in the Candoninae. For
some genera (e.g., Cypricercus) the testes form a spiral
around the inner edge of the shell rather than being
confined to the posterior area. The testes of Cyprididae
change into the vasa deferentia as they leave the epidermis and enter the body cavity. The vas deferens joins up
with the ejaculation ducts or Zenker’s organs (Fig. 9a).
These large organs are made up of longitudinal muscles
20. Ostracoda
in the shape of a tube on which are found radiating
bristles and are covered in a chitinous sheath. The alternate contraction and expansion of these paired organs
forces the sperm into the hemipenes during copulation.
Zenker’s organs lie in a nearly horizontal position in the
posterior part of the body cavity and are visible in many
live specimens. The hemipenes are very complex structurally. The ostracode must rotate them before copulation can take place (Kesling, 1957, 1965; Danielopol,
1969; McGregor and Kesling, 1969a, b). The testes and
ovaries in Cytheridae are found next to the intestine,
Zenker’s organs are absent in this family. There are no
males in the Darwinulidae. For further details on the
anatomy and physiology, the reader is referred to Hoff
(1942) and Kesling (1951a, 1965).
III. LIFE HISTORY
Ostracode eggs are constructed in a way that allows
them to withstand physical and chemical extremes. The
time of hatching may be soon after being laid or staggered over time indicating diapause. Once the egg is
hatched, freshwater ostracodes will go through eight
molt stages before becoming a mature adult. The length
of time the organism lives depends on the species.
A. Egg
The freshwater ostracode egg is a double-walled
sphere of chitin impregnated with calcium carbonate.
The space between the two spheres is occupied by a
fluid (Wohlgemuth, 1914). These two characteristics of
the egg allow it to withstand desiccation and freezing.
Ostracodes develop from eggs which may or may
not have been fertilized, depending on whether the
species is bisexual or asexual. The eggs are laid singly or
in masses, most often in the part of the habitat that will
ensure an oxygen-rich environment when the eggs hatch
(Wohlgemuth, 1914). Most ostracode eggs are white,
but some are green (Cypridopsis vidua) or bright orange (Cyprinotus incongruens, Potamocypris smaragdina). All North American freshwater ostracodes are
oviparous except several species of Darwinula and the
marine brackish water Cyprideis which are ovoviviparous. These taxa brood their eggs in the posterior of the
carapace from which the nauplii are released. Brood
pouches are more common in marine ostracodes.
B. Reproduction
Reproduction in freshwater ostracodes may be either sexual or asexual. Depending on the gender, the
ostracode has a pair of genital lobes or hemipenes.
817
Zissler (1969a, b, 1970) studied in detail the formation of sperm in Notodromas monacha. A single nucleus
is formed by fusing of caryomeres; it is characterized by
three granulated particles and by a double membranebounded region. The body then becomes spindle-shaped
which includes the nucleus and the “Nebenkern” derivatives. Further development of the spindle-shaped cells
takes place by the extension of two winglike structures,
“Flügelstruckturen,” on either side of the nucleus. The
nucleus and the winglike structures extend throughout
the sperm body, including the thread. The winglike
structures are interpreted as providing motility to the
sperm. Further differentiation of the sperm occurs between the testes and the vas deferens.
Lowndes (1935), following the research of other
authors publishing between 1854 and 1889, reassessed
the morphology and function of the sperm and concluded, “apparently the sperms are now functionless.”
Ostracode sperm have the dubious distinction of being
the longest of any sperm relative to the host animal
(Lowndes, 1935). Moore (1961) indicated that sperm
do serve a function because females only lay eggs after
copulation. However, Moore did not cite references to
support his claim. Havel et al. (1990a) demonstrated
that sperm have a function. Populations with males examined electrophoretically show genotypic frequencies
in close agreement with those expected in a randomly
mating sexual population.
Female-to-male sex ratios appear to vary depending on the species. Chaplin (1993) found that for
Candonocypris novaezelandiae there are two morphs.
For the “little brown morph” the sex ratio changes
from 2.8:1 to 6.9:1 over a 27-month period. The conclusion was that the sex ratios show an increase in the
abundance of parthenogenetic females. Dezfuli (1996)
reports a sex ratio for Cypria reptans of 2:1. Sex ratios
of modern North American ostracodes of 2:1 show a
female bias (Chaplin et al., 1994). These authors indicate there are no reports that male eggs are larger than
female eggs so there is no logical explanation for the
excess of females.
Based on earlier cytological studies, Bell (1982)
concluded that all ostracodes are diploid. White
(1973) indicated the diploid chromosome number
varies between 12 and 19. Chaplin et al. (1994) provide an up-to-date review of the sex mechanisms for
freshwater ostracodes. Although males are the heterogamic sex, some species have XO or XY sex
determination systems. A case of multi-X systems is
found in Cyprinotus incongruens where the females
have six pairs of X chromosomes and the males have
six X and a Y chromosome (Chaplin et al., 1994).
Further, they report that males of this species produce
two types of sperms, one with five and the other with
818
L. D. Delorme
10 chromosomes. Lécher et al. (1995) found some
parthenogenetic ostracodes have “supernumerary
chromosomes,” and that they have complex sex chromosome mechanisms with XO, XY, and multiple X’s
and Y’s. Three Canadian sexual ostracodes (Cypricercus splendida, C. deltoidea, and C.tincta) possess 22
chromosomes (Turgeon and Hebert, 1995). For C.
deltoidea and C. tincta meiotic pairing was seen. These
authors further indicate the asexuals, Cypricercus horridus, C. reticulatus, and C. fuscatus, had variable
chromosome counts and no evidence of meiotic pairing was observed.
Electrophoretic studies by Havel et al. (1990b)
show that 10 of 12 asexual ostracodes included clones
which were polyploid. They also found that clonal diversity varied among species. For instance, Cyprinotus
incongruens averaged 1.2 clones (1.3 clones; Havel and
Hebert, 1993), whereas Cypridopsis vidua had 8.3
clones (9 clones; Havel and Hebert, 1993), Cypricercus
reticulatus averaged 7.8 clones (Havel and Hebert,
1993) per pond. On a regional basis, high-diversity
species indicate a range of 18 – 80 clones, while low
diversity species show 1 – 7 clones (Havel and Hebert,
1993). Bell (1982) has indicated that asexuals are
more abundant in modern habitats, whereas, sexuals
are more abundant in habitats such as glacial lakes.
The study of ostracodes from various habitats from the
Canadian Arctic to the southern U.S.A. (Havel and
Hebert, 1993) to tropical habitats in Jamaica (Little
and Hebert, 1994) does not show an influence of
latitude on the prevalence of asexual vs sexual reproduction. Rossi et al. (1998) reported similar results
for Europe.
Hybridization has been documented between the
asexual Cyprinotus incongruens and C. glaucus (Turgeon and Hebert, 1994; Chaplin and Hebert, 1997).
Diploid asexual species when fertilized by a sexual
species produce an asexual triploid. Suspected hybridization between Cypricercus reticulatus and other
sexual cypricercids in Canada has been postulated
(Turgeon and Hebert, 1995). Theoretical models predict sexually reproducing species should have selective
advantage in unstable environments; however, a study
by Chaplin and Ayre (1989) does not indicate a complete association of sexually derived recruitment in unstable environments.
Darwinula is thought to be the oldest asexual ostracode. Schön et al. (1998) believe that, because of
this, accumulation of mutations should have occurred
between alleles “with lineages and between lineages”
through parthenogenesis. However, their data shows
the opposite; there is “no variability in the nuclear ITSI
region” despite analyzing individuals in the geographic
range between Finland and South Africa.
C. Embryology
Embryology of ostracodes has been studied by
Woltereck (1898), Schleip (1909), Müller-Cale (1913),
and Weygoldt (1960). The small size, double-walled
sphere, and textured outer surface of the egg does not
allow for direct observation of the cleavage process. It
is known, however, that after seven divisions, a 128celled blastula is developed (Kesling, 1951a). Differentiation into ectoderm and endoderm (entoderm of
Kesling) occurs after the eighth division.
D. Hatching and Seasonality
Most species begin hatching in the spring in the
temperate zone. The timing of this process is controlled
in part by temperature and geography (latitude).
Species which live in temporary (vernal) ponds have a
shorter life cycle. For example, Cypridopsis vidua develops in about one month (Kesling, 1951a) and during
several months for Cypria turneri (Strayer, 1988). Some
of these same species can also exist in permanent ponds
where several generations per year may develop. For
some species [e.g., Candona caudata; Delorme, 1978,
1982 (may belong to Fabaeformiscandona, see Meisch,
1996); not confirmed for North America, Cyprinotus
carolinensis McLay, 1978c], hatching is delayed and
spread over a much longer period. These apparent
“resting eggs” allow the animal to sustain itself in the
habitat in spite of stressful periods (such as anoxia and
pollution; Delorme, 1978) when the nauplii cannot
survive.
Eggs produced by some species can be either spring
or summer forms. The ostracodes hatched from eggs
laid in the spring do not tolerate warm summer temperatures. Ostracodes hatched from eggs laid in the
summer will live only in warm water and expire in cold
water (Schreiber, 1922). Martens et al. (1985) have
noted that eggs of the Australian Mytilocypris henricae
would not hatch below a certain temperature. Eggs of
freshwater ostracodes can withstand freezing (Kesling,
1951a; Sohn and Kornicker, 1979). As early as 1915,
Alm noted that temperatures affected the time of
hatching. Ephemeral ponds in which the sediments stay
frozen for up to 6 months during the winter produce
many nauplii from eggs in the spring. Angell and Hancock (1989) found that eggs of Heterocypris incongruens could withstand freezing and remain viable down
to 18°C when either wet or dry. Some authors have
hatched ostracode eggs from dry lacustrine sediments
kept in their laboratories for periods ranging from a
few years to many decades (Sars, 1901; Sharpe, 1918).
McLay (1978b) reported the eggs of Cypricercus reticulatus do not hatch until dried. Angell and Hancock
819
20. Ostracoda
(1989) found that eggs of Heterocypris incongruens remained viable after drying at 22 and 40°C. The eggs
did not remain viable if kept wet and heated to 40°C.
The eggs of some species (Cypridopsis vidua,
Cyprinotus incongruens, Physocypria sp., and Potamocypris sp.) can pass through the lower intestinal tract of
both wild and domestic ducks and remain viable (Proctor, 1964). Kornicker and Sohn (1971) reported that
ostracode eggs were still viable after being egested by
goldfish and swordtail fish. In an excellent paper detailing a meticulously carried out experiment, Rossi et al.
(1996) have shown that, during its life time, the same
individual of Heterocypris incongruens can produce diapausing eggs and eggs that hatch within a few days.
Angell and Hancock (1989) categorize eggs that hatch
within 10 days of being laid as being subitaneous.
Rossi et al. (1991) and Rozzi it al. (1991)have shown
the ratio of resting eggs depends on the genotype, temperature and photoperiod. Further they indicate the
percentage of dormant eggs increases with temperature
and daylight hours in a winter clone and decreases in
the summer clone. Also their studies show egg hatching
decreases at higher temperature, unaffected by photoperiod, in the winter clone and increases with both
temperature and daylight hours in the summer clone.
Rossi et al. (1996) found rather high number of first
hatchlings (neonates) at the end of the experiment for
the winter clone when they removed the algal mat to
count cluster of eggs. They suggest that bioturbation
may play a role in the hatching of eggs under field conditions. Rossi and Menozzi (1990) found from laboratory experiments that the dominant clone from the field
have a better chance of survival at low temperatures
and conversely the dominant summer clone survives
better at high temperature. This indicates that temperature is most likely responsible for seasonal cycles.
Most ostracodes are controlled seasonally in their
habitat. Research on seasonality has concentrated on
those species living in temporary ponds. For temporary
ponds in the temperate zone, early spring collections
made after the basin receives water yield few live ostracodes [Hoff, 1943a (Illinois), Ferguson, 1958a (South
Carolina)]. In these ponds a regular succession of juveniles can be observed until the pond dries, with some
species exhibiting at least two generations per year. A
temporary coastal habitat investigated by McLay
(1978a, b, c) has ostracodes from the fall through the
late spring. Ostracodes were absent through the summer because of habitat desiccation. Freezing had only a
marginal effect on a part of the habitat. McLay
(1978b) stated the maturation rate for Herpetocypris
reptans was 190 days, and 45 days for Cyprinotus
carolinensis. In permanent ponds, active species are absent for several months [Ferguson, 1944 (Missouri)].
Juveniles began appearing in early spring and continued until late spring, while adults were present until
late fall to early winter. This pattern was repeated for
the three species studied by Ferguson: Cypridopsis
vidua, Potamocypris smaragdina, and Physocypria
pustulosa. Ostracodes that live in ponds or ephemeral
bodies of water have a short life cycle. Martens et al.
(1985) noted that an Australian species took longer
(4 – 5 months) to reach sexual maturity in the winter
than in the summer (2 – 3.5 months). Candona subtriangulata, Cytherissa lacustris, and Darwinula stevensoni, that inhabit large lakes where the habitat is considered stable, will have a life cycle of 6 – 24 months
(McGregor, 1969; Delorme, 1978; Ranta, 1979). The
length of the life cycle depends on the species.
Danielopol (1980b) observed that the length of the life
cycle for five species ranged from 126 to 489 days, the
latter being for a hypogean ostracode.
E. Molting
During the eight molt stages, the appendages
change in size, shape, and function. A summary of these
developments is found in Table I (after Kesling, 1951a).
Przibram (1931) determined that crustaceans doubled their weight from one molt to the next. He further
concluded that because weight varies directly with volume, that volume would also increase by a factor of
two. This led him to believe that a given linear dimension would increase by about the cube root of two
(1.26) for each molt. Based on this hypothesis,
Kesling (1953) devised a circular slide rule which allows
one to determine the variability of any linear dimension,
area, or volume of any ostracode molt stage and adult.
This slide rule is useful in determining which molt stage
(shell or appendage) one is examining. Being able to
identify an ostracode as an instar saves time because
most instars cannot be identified to the species level.
TABLE I The Development of Appendages with Each Instar
of the Freshwater Ostracodea
Instar
1
2
3
4
5
6
7
8
9
Antenna I
Antenna II
Mandible
Maxilla
Furca
Leg I
Leg II
Leg III
Ovaries
Testes
a
() indicates the anlagen of a structure.
820
L. D. Delorme
F. Competition, Co-existence, and Survival
The study of competition, coexistence, and survival
by McLay (1978a – c) showed that, in a small temporary
puddle, “competition is mediated via density-dependant
mortality and maturation rates.” He found that Herpetocypris reptans was a better competitor because of its
longer maturation time (4419 degree days vs Cyprinotus
with 2034 degree days) and could better withstand
crowding and a shortage of food. His model shows that
Herpetocypris was dominant over Cyprinotus carolinensis by a factor 7:1. He also found that competitive
exclusion of Cyprinotus by Herpetocypris was prevented
by periodic freezing. Freezing was also found to harm
the population growth of Herpetocypris more than
Cyprinotus because of the longer time required for maturation. Oertli (1995) studied the density of invertebrate
of three substrates of a manmade pond. He found that
life cycles and substrate controlled density fluctuations.
High densities in the summer are related to life cycle and
newly hatched individuals. He called the summer effect
the “reproduction effect” with high mortality occurring
in the winter and spring. For Cypridopsis vidua the
highest densities occurred on Chara and Typha, with the
lowest densities on terrestrial leaf litter. Rieradevall and
Roca (1995) found that substrate and organic matter are
important factors controlling the distribution of ostracode species in a karstic spanish lake. They collected
high densities in the fall and spring.
IV. DISTRIBUTION
Ostracodes are found in nearly every conceivable
aquatic habitat. These range from temporary and permanent ponds, lakes, intermittent, and permanent
streams to ditches and irrigation canals, and fens. Caves
and the interstices of aquifers are additional habitats.
Some (referred to as terrestrial ostracodes) are found in
moist organic mats of fens and in axial cups of certain
plants such as bromeliads. North American freshwater
and brackish water ostracodes thrive in a full range of
salinities from 10 to 74,800 mg / L [high sulfate, low
chloride waters] (personal files). Brackish water forms
are common in high sulfate waters (Limnocythere staplini), as well as in chloride waters mixed with freshwater (Cytheromorpha fuscata) in inland lakes (Neale and
Delorme, 1985). De Deckker (1981) lists an Australian
species found in waters having salinitics greater than
170,000 mg / L. Cyprideis, also high salinity species, are
common in southern coastal areas of North America
(Benson, 1959), England and the Netherlands (van
Harten, 1975; Sywula et al., 1995). Jahn et al. (1996)
have found that Cyprideis torosa can oxidize hydrogen
sulfide to nontoxic thiosulfate and sulfite and rid itself
of the oxidation products quickly.
A. Benthic Swimmers
Ostracodes are benthic organisms and are only
rarely found in the plankton. They are divided into
swimmers and nonswimmers. Most often they swim
between aquatic plants. If they are disturbed while
swimming, they often lose control and spiral down to
the substrate. A few species of the genera Cypria and
Physocypria are known to swim up into the water column, as they have been recovered from suspended sediment traps. Another swimmer, Notodromas monacha,
has a peculiar habit of turning upside down when approaching the water surface. The venter of this species
is flat, enabling the organism to adhere and hang beneath the water surface by surface tension. Marmonier
et al. (1994) indicate that plant dwelling species, such
as Cyclocypris ovum and Cypridopsis vidua, are more
likely to be spherical in shape.
B. Benthic Nonswimmers
The nonswimmers, or true benthic forms, use their
well-developed antennae (without setae) and first thoracic legs (previously called the maxillae) to crawl on the
sediment – water interface. Some species, such as the limnocytherids, are infaunal, crawling beneath the sediment – water interface between the sediment particles (L.
Delorme, personal observations; Benzie, 1989). Ostracodes live in the vents of warm and cold-water springs
where groundwater is discharging (Forester, 1991;
McKillop et al., 1992). Särkkä et al. (1997) has reported
Potamocypris pallida, Candona candida, and Candona
reducta made up 21% of a spring meiofauna affected by
road de-icing salt and gravel extraction. Ostracodes may
live in the fractures of bedrock adjacent to streams,
lakes, and pools in caves.
Hypogean (subterranean) ostracodes are being recovered more frequently from aquifers. These blind animals live in the interstices of sand aquifers. They are
recovered by pumping the aquifer or trenching through
it (Danielopol, 1980a). Marmonier et al. (1994) have
shown that interstitial ostracodes have a geometric
shape (triangular or trapezoidal) which is most suited
to its habitat. The same can be said for most limnocytherids which live just below the sediment – water
interface, crawling between the sediment interstices
(personal observations). Ward et al. (1994) observed
the highest concentration of hypogean ostracodes in a
well close to the main channel of the Flathead River,
Montana, and in wells farthest on the floodplain but
near the Whitefish River. Rogulj et al. (1994) indicate
20. Ostracoda
those hypogean ostracodes (Fabaeformiscandona
wegelini) that live in shallow interstitial habits are
closely linked to running water rich in organic matter.
This concept is supported by observations of Rouch
and Danielopol (1997) and Marmonier and Creuzé des
Châtelliers (1991). Creuzé des Châtelliers and Marmonier (1993) determined there were no meaningful
correlations between abundance of ostracodes and particulate organic matter, alkalinity, calcium, or oxygen
content of the interstitial water. Creuzé des Châtelliers
and Marmonier (1990) found the interstitial fauna
above and below a river riffle shows differing ostracode
assemblages and abundances. Above the riffle, where
the river bed was degrading, the water velocity was
greater and the number of species and specimens was
less than below the riffle. Below the riffle the river bed
was aggrading and the velocity of the water was less,
allowing more species to live in this area both in composition and numbers. Those living in deeper aquifers,
prefer a more stable environment with less organic
matter (Rogulj et al., 1994). Along the length (320 km)
of the Rhône River, Marmonier and Creuzé des Châtelliers (1992) found three different species assemblages
tied to the ecological requirements of the member
species. Two groups were restricted to the upper and
lower ends of the river respectively and one was common to the whole length of the river.
821
B. Salinity and Solute Composition
Salinity and solute composition are of importance
to freshwater ostracodes. Delorme (1989) has shown
preliminary salinity tolerance distributions for some 43
species studied from Canadian habitats (Fig. 13). The
sequence of species shown for salinity tolerance shows
a gradual increase in the mean salinity value. Furthermore, the range, as shown by minimum and maximum
values of salinity tolerance, is a function of the species.
For example, Cypridopsis vidua, is a moderately salinetolerant species able to survive in a very broad range of
salt concentration. On the other hand, species such as
Candona subtriangulata and Cytheromorpha fuscata
are much more restricted in their chemical, and thus
geographic range. The low end of the range, or
pristine, low-salinity (SO4) waters are inhabited by
such species as Candona subtriangulata (23 – 93 mg / L),
V. CHEMICAL HABITATS
A. Chemical Water Types
Physical and chemical characteristics of the habitat
are important factors in ostracode distribution and
abundance. From Canadian habitat-waters, four chemical types are defined based on the predominant equivalent anion or oxyanion. These are bicarbonate, sulfate,
chloride, and carbonate (personal files). A high percentage of genera (61%) are found only in bicarbonate waters. Bicarbonate, after being converted to carbonate, is
a primary constituent of the shell. Only 9% of the
known genera are found living in sulfate-rich waters.
Several genera (30%) have species which have a preference for either of the two types of water. Chloride-dependent species, such as Cytheromorpha fuscata and
species of Cyprideis, are more properly tolerant of marine brackish water. C. fuscata are occasionally found
in inland lakes (Neale and Delorme, 1985) where brine
seepages occur. Such species as Candona rectangulata,
Cyprinotus salinus, and Cypris bispinosa are found in
lake and pond habitats that may receive sea spray.
These species are by no means restricted to chlorideenhanced waters.
FIGURE 13
Total dissolved solids range for some Canadian freshwater ostracodes. The solid line represents the mean, the bars
represent the minimum and maximum values. [Modified from
Delorme, 1989.]
822
L. D. Delorme
Limnocythere verrucosa (22 – 255 mg / L) from the
Great Lakes area, and Candona protzi (may belong to
Fabaeformiscandona, see Meisch, 1996, not confirmed
for North America) (56 – 115 mg / L) of the high arctic.
The high end of the range or high-salinity water bodies
are inhabited by such species as Limnocythere ceriotuberosa (74 – 2780 mg / L) and L. staplini (122 –
74,840 mg / L). De Deckker (1983a) reported there are
14 North American ostracode species (one marine) that
live in athalassic saline waters. Species such as
Cyprinotus glaucus may be found in lakes where there
is groundwater discharge of saline waters of intermediate flow systems (Delorme, 1972).
Forester (1983, 1986) and Forester and Brouwers
(1985) have emphasized the importance of water composition over salinity. Examination of lake-water chemistry in North America, where Limnocythere sappaensis and L. staplini live, reveals that solute composition
and not salinity is most important for their existence
(Forester, 1983; Delorme personal files). Forester found
that L. sappaensis lives in water enriched in
Na –HCO3 –CO32 and depleted in Ca2 . L. staplini
lives in water enriched in various combinations of
Na2 –Mg2 –Ca2 –SO42 –Cland depleted in HCO3.
Forester (1987) and De Deckker and Forester (1988)
have expanded on the relationship between ostracode
occurrences and major dissolved ion chemistry and
salinity. They further suggested that relationships between species diversity, carbonate saturation, and relative abundance of dissolved calcium and carbonate ions
exist, based on limited data from the United States.
Most ostracode species may be placed in one of six or
seven hydrochemical groups. The groups have welldefined boundaries based on carbonate saturation, major dissolved ion composition, and salinity.
FIGURE 14
pH range for some Canadian freshwater ostracodes.
The solid line represents the mean, the bars represent the minimum
and maximum values. [Modified from Delorme 1989.]
C. Hydrogen Ion Concentration
The preservation of the calcareous exoskeleton for
live ostracodes suggests that both the pH of the habitat
must be circum-neutral and that a ready source of
CaCO3 must be available. Delorme (1989) indicated the
“average” pH of Canadian aquatic habitats for ostracodes is between 7 and 9.2 (Fig. 14). For carbonate-enriched lakes, common to groundwater discharge areas
of the United States and Mexico, pH values are typically in the range of 9.6 – 10.5 (R. M. Forester, personal
communication). The species has an individualized pH
range. Several of the species have a very broad pH
range such as Cypria ophtalmica (5.2 – 13), Candona
candida (5.4 – 13), Cypridopsis vidua (5.2 – 12), Cyclocypris sharpei (5.2 – 11), and Cyclocypris ampla (5 – 12).
Cytheromorpha fuscata with its limited distribution in
the Manitoba Lakes has a very restrictive range of 8.3
to 8.7 in inland lakes. This may be an artifact of the
lakes from which the species was collected. The majority of ostracodes have a larger pH range than C. fuscata. When the epidermal lining covering the shell is
ruptured and the water and sediments are poorly
buffered, the calcareous shell will dissolve, destroying
the animal. Larsen et al. (1996) used a number of statistical method to investigate the relationship between
invertebrates to pH in Norwegian river systems. They
indicate that a significant response occurs in the abundance of species when there is an increase in pH.
D. Dissolved Oxygen
Dissolved oxygen in the aquatic habitat is an important requirement for survival. The mean requisite
20. Ostracoda
823
oxygen content below its lower tolerance limit of
2.8 mg / L. Its survival was accomplished by having a
short life cycle allowing it to produce eggs before the
onset of anoxia. This survival mechanism has been in
place for some time in Lake Erie as indicated by the
presence of fossil shells in the lake sediments (Delorme,
1982). Contrary to C. caudata (may belong to Fabaeformiscandona; see Meisch, 1996, not confirmed for
North America), C. subtriangulata and Cytherissa lacustris have become locally extinct in the central basin
of Lake Erie because they have a 1-year life cycle and
cannot reach sexual maturity as they require a minimum dissolved oxygen content of 5.6 and 3.0 mg / L.
Cytherissa lacustris is making a comeback with a reduction in anoxia. Newrkla (1985) found C. lacustris
could survive at 1 mg O2 /L in the laboratory on a glass
substrate and minimal food (20°C and 20-h exposure).
Nine of the 43 species listed in Figure 15 can tolerate
near-zero dissolved oxygen (below the detection limit
of the Winkler azide method, APHA, 1965) for short
periods. Candona decora, C. candida, Cyclocypris ampla, C. sharpei, and Cypria ophtalmica have been recovered from water of zero dissolved oxygen from a
small river in early December (L. D. Delorme, unpublished data). The river had been covered by 1 ft of ice,
suggesting these waters had been anoxic for some time.
Fox and Taylor (1954, 1955) have found by experimentation that some ostracodes survive longer at oxygen levels below air saturation of 21%.
E. Carbonate
FIGURE 15 Dissolved oxygen range for some Canadian freshwater
ostracodes. The solid line represents the mean, the bars represent the
minimum and maximum values. [Modified from Delorme, 1991.]
for dissolved oxygen by ostracodes falls within a very
narrow margin of 7.3 to 9.5 mg / L (Fig. 15). In general
the concentration of dissolved oxygen in the water is
broad. Candona subtriangulata has the highest minimum oxygen requirement of 5.6 mg / L. Most surprising in Figure 15 is the high number of species (34 out
of 43, in Canadian habitats, L. D. Delorme, unpublished data) which can tolerate low dissolved oxygen
concentrations. Those species which require a minimum 3 mg / L dissolved oxygen in the water are
Cytherissa lacustris, Cytheromorpha fuscata, Ilyocypris
gibba, Limnocythere ceriotuberosa, L. herricki, L.
itasca, L. verrucosa, and Potamocypris variegata. Delorme (1978) found that Candona caudata (may belong
to Fabaeformiscandona, see Meisch, 1996; not confirmed for North America) could survive in Lake Erie
even though the hypolimnion has summer dissolved
The effect of excess calcium carbonate and anoxia
on Cypridopsis vidua was investigated by Reyment and
Brännström (1962). Their results indicated the morphology of the shell is adversely affected (smaller shell
dimensions) by excess calcium carbonate and anoxia in
the environment. They reiterate the common belief that
the major contributor to the size and shape of an ostracode is the environment, with a secondary contributor
being the genome of the animal. Van Harten (1975)
has shown the effect of salinity on shell size. The
largest mean size of Cyprideis torosa was found in
nearly freshwater, whereas the smallest mean size was
found in waters where the salinity was very high.
VI. PHYSICAL HABITAT
A. Temperature
The water temperature of the aquatic habitat plays
an important role in the habitat, seasonal, and geographical distribution of ostracodes. This is not to
824
L. D. Delorme
say that other factors such as availability of food, solute
composition, turbidity, and energy levels are not important. The temperatures in shallow-water habitats can
range from 0 to over 30°C, depending on the latitude
and altitude. Species living in this niche must have the
ability to tolerate this broad range in one or more life
stages. Such species as Candona acutula, C. candida, C.
ohioensis, Cyclocypris ampla, C. sharpei, Cypria ophtalmica, and Limnocythere staplini exhibit such a range
for bottom water temperature (Fig. 16). The exception
to the above are shallow ground water fed streams
which may have a nearly constant temperature.
Candona subtriangulata has a low mean value of
5.5°C, with a range of 2.6 to 19.2°C (Delorme, 1978).
This species is common in Lakes Superior, Huron, and
Ontario at considerable depth, where the bottom-water
temperature does not vary much.
Korschelt (1915) ran several experiments with
Cypridopsis vidua and Cypria ophtalmica, in which the
species were frozen in ice for several hours. The ice was
then allowed to melt; the majority of the specimens
survived the freezing experiment.
At the other temperature extreme, Cypris balnearia
has been described from thermal springs at temperatures of 45 to 50.5°C. Wickstrom and Castenholz
(1973) have recovered Potamocypris from an
algal – bacterial substrate of a hot spring, in Oregon, at
temperatures ranging between 30 and 54°C. Chlamydotheca arcuata has been found in warm springs (Utah,
Nevada, Arizona, and Mexico) where water temperature varies between 24 and 39°C (Forester, 1991).
Mean annual air temperature assists in explaining
the geographic distribution of ostracodes in habitats
shallower than 25 m. From Figure 17, 20 out of 43
FIGURE 16 Bottom water or habitat temperature range (°C) for
some Canadian freshwater ostracodes. The solid line represents the
mean, the bars represent the minimum and maximum values.
[Modified from Delorme, 1991.]
FIGURE 17 Mean annual temperature range (°C) for some
Canadian freshwater ostracodes. The solid line represents the mean,
the bars represent the minimum and maximum values. [Modified
from Delorme, 1991.]
20. Ostracoda
species can live in aquatic habitats north of latitude
60°. The minimum values of the range for mean annual
air temperature for these species habitats are between
8.5 and 11°C. The species restricted to the arctic
and low mean annual air temperatures are Candona
paralapponica, C. protzi (may belong to Fabaeformiscandona; see Meisch, 1996, not confirmed for North
America), C. anceps, C. pedata, C. mülleri, C.
ikpikpukensis, Cyclocypris globosa, Limnocythere liporeticulata, and Tonnacypris glacialis . The habitats
for those species living in the midlatitudes have a mean
annual air temperature greater than 1.5°C.
B. Lakes, Ponds, and Streams
Most freshwater ostracode species may be found in
more than one habitat (Fig. 18). The pond – lake habitats
825
can be viewed as a continuum from shallow to deep
water. Some ponds are temporary in nature and, therefore, harbor a specific faunal association such as Candona renoensis, Cypricercus deltoidea, Megalocypris
alba, and Cyclocypris laevis (Delorme, 1989; King et al.,
1996; Wiggins et al., 1980). At the opposite end of the
scale are those species adapted or restricted to living
in permanent lake habitats (Candona subtriangulata,
Cytherissa lacustris, and Cytheromorpha fuscata). A
faunal mix may occur between pond-lake and stream
habitats. There may be a specific biotope developed at
the mouth of a stream or river where it enters the lake or
pond. The delta area is characterized by a combination
of flowing and standing water. Many ostracode species
are adapted but not restricted to living in this biotope
such as Candona acuta, Cyclocypris ampla, C. sharpei,
Ilyocypris bradyi, I. gibba and Potamocypris variegata.
Large lakes have bottom currents which accommodate
species such as Candona caudata (may belong to Fabaeformiscandona; see Meisch, 1996, not confirmed for
North America) and C. acuta. None of the species listed
by Delorme (1989) are completely restricted to flowing
water. Those which are considered as lotic (fluvial)
species are Candona acuta, Cypria obesa, Ilyocypris
bradyi, I. gibba, and Potamocypris variegata. A special
case is the fen. Up to 10 species have been recovered
from fen vegetation at various levels down to 20 cm and
varying levels of dissolved oxygen and redox potential
(Douglas and Healy, 1991).
C. Bromeliads
Axial cups are formed by the spiral overlap of
bromeliad leaves. These cups collect rain water and become inhabited by many kinds of invertebrates.
Tressler (1956) described Metacypris maracoensis,
recovered from the axial cups of Florida bromeliads.
Ostracodes are more commonly found associated with
bromeliads in Central and South America, and the
Caribbean Islands (Little and Hebert, 1996).
D. Cave and Subterranean Habitats
FIGURE 18 Percentage probability of finding a Canadian freshwater
ostracode in a lake, pond, or stream. [Modified from Delorme, 1989.]
Ostracodes have been recovered from several caves.
Klie (1931) reported Entocythere donnaldsonensis commensal on the crayfish Cambarus from the Donnelson
Cave of Indiana. From the Marengo Cave of Indiana,
Klie named two new species:Candona marengoensis
and C. jeanneli. Hart and Hobbs (1961) have described
eight new troglobitic cave species from the eastern
United States. Walton and Hobbs (1959) have also described troglobitic ostracodes from Florida caves.
Pools in deep sinkholes in limestone terrain
(cenotes) of Yucatan, Mexico have a rich and varied
826
L. D. Delorme
ostracode fauna. Furtos (1936a) described 23 species
from these habitats.
More research has been carried out on hypogean
ostracodes. Cavernocypris wardi has been recovered
from spring and interstitial water of the Flathead River,
Montana (Ward et al., 1994), from the South Platt
River, Colorado (Danielopol et al., 1994; Marmonier
et al., 1989; Marmonier and Ward, 1990 (Candona
candida, Cavernocypris wardi, Cypria sp., Eucypris sp.,
Fabaeformiscandona pennaki, F. wegelini, Ilyocypris
sp., Nannocandona faba, Potamocypris sp., Pseudocandona cf. P. albicans, Strandesia canadensis), and from
springs in Wyoming, Nevada, and Colorado (Forester,
1991). Cavernocypris subterranea (common in Europe)
has been found from a spring near Brush Creek, Idaho
(Külköylüoĝlu and Vinyard, 1998). Not all ostracodes
found near spring discharges are hypogean (Forester,
1991; McKillop et al., 1992; Ward et al., 1994).
E. Sediment – Water Interface, Particle Size, Food
The distribution of ostracodes at the sediment –
water interface is a function of the availability of food,
whether the substrate surface is clearly or poorly
defined, the particle size distribution of the top centimeters of the substrate, as well as the time of year.
The distribution of ostracodes is patchy. Where there is
organic detritus, ostracodes will be abundant, particularly in shallow water. The presence of abundant floating organic debris just above an ill-defined sediment –
water interface, discourages ostracodes from living in
this part of the habitat. As detritivores and herbivores,
ostracodes require a ready source of particulate organic
matter that they can sweep into the mouth region.
They also use their mandibular teeth to rasp or gnaw
off small particles from large organic particles, living
plants, or algae. On predominantly mineral substrates,
ostracode densities will be low. As the sediment – water
interface becomes deeper and further from the shoreline the number of species decrease. In this niche, the
only food available will be the resistant cellulose raining down through the water column, bacteria, and
fungi. In the deep troughs located in the eastern end of
Lake Superior, only Candona subtriangulata has been
recovered from depths of 45 – 305 m. Tressler (1957)
identified Candona decora at a depth of 600 m from
Great Slave Lake, Canada.
For many limnocytherids, especially Limnocythere
ceriotuberosa, and L. staplini, the particle size of the
substrate is very important. These ostracodes crawl
through the interstices of very fine sand (0.0625 mm)
to coarse sand (1.0 mm). In these niches, there must be
sufficient food and oxygen for survival. Good porosity
and permeability of these sediments allows a freeflow
of oxygenated waters. Most clay-sized, organic-rich
sediments are anoxic and reducing and not suitable for
ostracodes to live in the interstices.
VII. PHYSIOLOGICAL AND
MORPHOLOGICAL ADAPTATIONS
Ostracodes live in a wide variety of habitats, and
within a given habitat physical and chemical conditions
fluctuate. Physiological adaptations can be seen for
some of these changes.
A. Eggs
Dispersal of freshwater ostracode eggs is thought
to be passive (Neale and Delorme, 1985; Peck, 1994;
Sywula et al., 1995; Little and Hebert, 1996;
Malmquist et al., 1997) either in the gut or mud on the
feet of birds, particularly for species living in the littoral habitat. Eggs that pass through the gut of fish and
remain viable assist in passive dispersal (Kornicker and
Sohn, 1971). For sampling sites that were up to
1000 km apart, Chaplin and Ayre (1997) found no
evidence that stream flow was a “mediator of short or
long distance gene flow in (the large green morph of)
Candonopsis novaezelandiae.”
B. Resting Stage and Torpidity
The primary resting stage for freshwater ostracodes is the egg. The structure of the egg permits the
yolk to survive both desiccation and freezing. The
other adaptation for surviving harsh conditions in ostracodes is torpidity (Barclay, 1966; Delorme and Donald, 1969; McLay, 1978a; Horne, 1993). Candona
rawsoni has been recovered in the seventh instar from
frozen sediments of a pond which had gone dry the
previous fall. Placing the sediments in water at room
temperature caused rejuvenation of the seventh instar
candonids. Survival rate was a function of the moisture
content and temperature of the sediments holding the
instar at the time of rejuvenation (Delorme and Donald, 1969). The major difference was that sediment
containing the seventh instar could have a moisture
content as low as 4 – 5%. In the fall or spring, hatching
of the eggs and development of the nauplii from the
seventh instar may occur. In vernal ponds, torpidity allows sufficient time for the species to become sexually
mature and lay eggs. In this way, the species is propagated in temporary bodies of water (Wiggins et al.,
1980). The development of the torpid state allows
ostracodes to survive the drying of vernal ponds.
Horne (1993) found a similar situation for Candona
20. Ostracoda
patzcuaro, collected from playa lakes of the south
plains of Texas.
C. Coloration
Coloration of ostracodes is a function of the pigment bodies located in epidermal cells of both the outer
and inner layer. Ostracodes such as Cypridopsis,
Cypris, Cypricercus, Eucypris, Herpetocypris, Potamocypris, and Tonnacypris are normally green, although
Cypridopsis vidua and Cypricercus may be found in
shades of brown and purple. These genera are all
prevalent in littoral areas of lakes and in ponds where
there is abundant vegetation. Green (1962) investigated
the green coloration of Eucypris virens and found it to
be a bile pigment, biladienes, derived from blue-green
algae. Other species, such as Cypricercus horridus and
Megalocypris alba, are purplish to reddish-brown in
color which allows them to blend better with an organic substratum. Mbahinzireki et al. (1991) found
that marked coloration increased predation risk by fish.
The majority of the other species are a dirty white to
yellowish-buff color which blends in with the mineral
substrata. Candonids, cyprinotids, and limnocytherids
fall into the last category.
D. Appendages
The first and second antennae are modified in
swimming ostracodes. Finely feathered swimming setae
on the first antennae are in constant motion and offer
resistance to the water when the ostracode is swimming. Setae on the second antennae are used in a rowing motion. When the swimming setae are absent, the
second antennae are used exclusively for walking and
climbing.
In the male of some species, the maxillae (formerly
referred to the first thoracic leg) is modified as a prehensile palp. This changes the appendage from a feeding function to a sexual function.
VIII. BEHAVIORAL ECOLOGY
Experiments were conducted by Towle (1900) on
heliotropism using Cypridopsis vidua. She determined
that ostracodes initially moved toward the light and
then reversed directions.
Ostracodes that swim maintain a sustained motion. The swimming setae of the first antennae used in
propelling the animal forward must be in constant motion or the animal will sink to the substrate. Observation of the swimming motion, using a binocular microscope, shows the swimming setae moving in a pattern
827
similar to a hand-held egg beater directed upward. Because swimming is energetically quite expensive, ostracodes can only swim short distances (usually between
adjacent plants).
The entry of a stimulus from a cyprinid fish tank
into a microcosm of Chara fragilis (Uiblein et al. 1994)
changed the behaviour pattern. The result was that
Cypridopsis vidua responded by spending more time
among the plant stems and leaves.
IX. FORAGING RELATIONSHIPS
The diet of most ostracodes is restricted to algae
(phytophilic) and organic detritus. Klugh (1927) reported certain algae as being part of the food supply.
Kesling (1951a) indicated diatoms are a part of the ostracode diet as did Strayer (1985) for Cypria turneri.
Grant et al. (1983) reported feeding Cyprinotus carolinensis the blue-green algae Nostoc sp. Campbell (1995)
calculated that tree pollen made up 8% and organic
material 42% of the diet for Australocypris insularis.
Tabacchi and Marmonier (1994) used Chara with alterable amounts of periphyton covering the stems as
did Roca et al. (1993). When feeding, organic particles
are swept into the mouth using the maxillae. The
mandibular teeth grind large organic particles into a
smaller size before they enter the mouth.
Although ostracodes are predominantly herbivores
and detritivores, a few have shown carnivorous characteristics (Johansen, 1912). Ostracodes have been observed attacking and eating the soft tissue of certain
snails (Deschiens et al., 1953; Deschiens, 1954; Sohn
and Kornicker, 1975). Some species (Cypridopsis
hartwigi, C. vidua, Cypretta kawatai, and Cyprinotus
incongruens) are known from experiments, by the
above authors, to eat soft tissue of snails, such as the
mantle and antennae, and to crawl into the respiratory
organs. Have (1993) used Paramecium sp. as a source
of food by Eucypris sp. in experiments on effects of
area and patchiness on species richness. Campbell
(1995) provides detailed information on the preying
habits of Australocypris insularis. These large ostracodes feed on smaller ostracodes, Diacypris compacta
and D. dietzi. She also found this large species grouped
around drowned bee carcasses, presumably feeding on
them, with terrestrial invertebrate detritus making up
8% and insects 3% of the diet.
The trophic position of most ostracodes is that of a
herbivore and detritivore. This group of benthic animals scavenges and consumes organic matter primarily
from the sediment – water interface. Some of this food
is fresh, while the remainder is in the process of decaying. During the feeding process, nutrients and the more
828
L. D. Delorme
resistant cellulose and silica (diatom frustules) are
moved back into the food chain.
Ostracodes are, in turn, consumed by higher animals. Predators include bottom dwelling fish. There are
several references to the presence of ostracodes in the
gut of fishes. Bigelow (1924), Harding (1962), and Ferguson (1964) found ostracodes in the stomachs of
suckers and trout. Stomach contents of brook stickleback contained Cyclocypris ampla, that of yellow
perch had Candona ohioensis, and bullheads had
Cypridopsis vidua and Physocypria globula (L. D.
Delorme, unpublished data). Griffiths et al. (1993) report the presence of Ostracoda in the guts of arctic
charr, Salvelinus alpinus, and the three-spine stickleback, Gasterosteus aculeatus (Dezfuli, 1996). Bataille
and Baldassarre (1993) investigated three prairie potholes (near the Delta Waterfowl Research Station,
Manitoba) as to the availability of aquatic organisms
(including ostracodes) as high protein food for ducks
such as mallards and canvasbacks. They used activity
traps to capture nektonic organisms during a 24-h period. The sample was then filtered through a #35 U.S.
Standard sieve (500 m). It should be noted that this
sieve size would lose most ostracodes except Cypris pubera. Batzer et al. (1993) investigated the feeding habits
of mallards and Green-Winged Teals on aquatic invertebrates. The contents of esophagi and habitat locations were studied using D-framed sweep nets (30-cm
width, 1-mm mesh size) for nektonic and epiphytic invertebrates. For sediment sampling of the habitat, a
bilge-pump intake was placed on the wetland bottom
and four pumping repetitions of water and sediment
were passed through a 0.3-mm sieve. This sieve size
would lose most ostracodes. Ostracodes and other invertebrates were each consumed by only 3 or 4 of the
19 mallards. The relative spacing of bill lamellae of
specific duck species influenced their selection of different orders of invertebrates (Nudds and Bowlby, 1984;
Batzer et al., 1993).
Green (1954) noted the oligochaete Chaetogaster
diaphanus eats ostracodes as do cyclopoid copepods
(Fryer, 1957) and the tanypodine midge (Roback,
1969). Benzie (1989) indicated “a negative association” between a predatory mite and Herpetocypris reptans. Swüste et al. (1973) showed experimentally that
the phantom midge Chaoborus flavicans released
Cypria immediately after capturing it. This suggests
that this ostracode either tastes bad or releases a noxious chemical. Lancaster and Robertson (1995) looked
at the preying habits of the polyphagous net-spinning
caddisfly (Plectrocnemia conspersa) and the alderfly
(Sialis fuliginosa). They found both flies had very high
densities of ostracodes in their guts, probably because
both groups are very abundant in leaf litter of slow
moving streams. The introduction of fish into a pond,
where larger predators have been absent, quickly reduces the invertebrate populations. Martens and de
Moor (1995) found the invertebrate populations that
lived in a large predator free habitat were adapted to
living in such an environment. The consequence is
these habitats will not be repopulated with the same invertebrate populations.
X. POPULATION REGULATION
A. Suitability of Habitat
Abundance of ostracodes at the sediment – water
interface is a function of substrates, availability of
food, season, and depth of water. Strayer (1985) has
provided a useful and current summary of abundance
and biomass measurements of ostracodes in lake environments. Bechara (1996) has indicated floodplain
lakes with high floating plant biomass had a decrease
in biomass during high-water periods, Ostracoda and
Oligochaeta were dominant. The presence of food
clearly has an impact on species richness and abundance. Long periods of warm temperatures down to
the bottom of the euphotic zone favor ostracode production. The number of species and specimens recovered from the profundal sediment – water interface is
small. For example, the ostracode assemblage in the
deep troughs (190 – 365 m) of eastern Lake Superior is
limited to Candona subtriangulata. At these depths,
during the last week in May and the first week in June,
the bottom water temperature varied between 2.6 and
4.0°C.
Based on Canadian habitats sampled for ostracodes, a maximum of nine genera have been counted
from one sample. In these samples, between 13 and 19
species were represented with a maximum of nine
species being alive at the time of collection (L. D.
Delorme, unpublished data). Rouch and Danielopol
(1997) defined high species diversity as “species collections of more than 25 species within an inventoried
area, at the local scale (i.e., between 1 and 100-m
length), low diversity is 1 – 5 species and medium level
diversity is 6 – 24 species.” King et al. (1996) found
species richness varied up to 13 species per pond. Baltanás (1992) attempted to make a comparison between
samples using species numbers referred to a fixed sampled area. He evaluated three methods for estimating
species richness (the number of individual species per
sample) using computer simulations. These were: Stout
and Vandermeer method based on the species – area relation; Cohen’s method which assumes the log-normal
distribution of species abundances; and the jackknife
20. Ostracoda
method, a nonparametric procedure with no distributional assumptions. The author found the estimators
tested showed a negative bias, with the second-order
jackknife method giving the best results with the highest accuracy and with the best precision.
B. Toxicity of Herbicides and Pesticides
Sanders (1970) tested Cypridopsis vidua and five
other crustaceans against 16 herbicides. Irritability and
excitability were the first noticeable reactions. Cypridopsis vidua could only be tested at low concentrations
because the animal would close its shell when it received the stimulus. This species was the second most
sensitive to the herbicides. Landis et al. (1994) found a
similar pattern when testing ecological risk assessment
of Cyprinotus sp. and other organism in a microcosm
against the water soluble fraction of jet fuel A. Dieter
et al. (1996) tested several invertebrates in microcosms
at the edge of lakes for toxicity to insecticides. They
found ostracodes were resistant to phorate. Takamura
and Yasuno (1986) found that ostracodes in rice fields
decreased in numbers with a treatment of pesticide
mixture of propoxur, thiobencarb, and simetryne.
There were similar decreases in ostracodes with a treatment of bentazone. In another rice field, they used a
mixture of methomyl, kasugamycin-hydrochloride,
neo-asozin, thiram, and benomyl with some decrease in
ostracode fauna.
C. Ultraviolet-B Radiation
Hurtubise et al. (1998) subjected a number of invertebrates to ultraviolet-B radiation. They found
that Cyprintous incongruens was highly tolerant at a
water depth of 4 cm when the UV-B irradiance was
56.1 W / cm. They also contend the carapace protected the animal. Further that dissolved carbon (DOC)
attenuated the irradiance.
D. Parasitism
More studies are now documenting the role freshwater ostracodes play in parasitism. Dezfuli (1996) has
detailed the effects of the helminth Neoechinorhynchus
rutili on the ostracode Cypris reptans. Between 8.6 and
14.5% of this species were infected, with up to three
larvae found in one host (up to six worms; see references in Zelmer and Esch, 1998). The effect of this parasitism is a reduction in the number of eggs contained
in females, 103 as compared to 207 in uninfected females. Cypridopsis vidua has been found to be the intermediate host of Neoechinorhynchus prolixus (B. B.
Nickol, University of Nebraska, personal communica-
829
tion). Cypridopsis sp. acts as a second intermediate
host for the parasite Halipegus occidualis (Zelmer and
Esch, 1998). For the complete cycle, eggs from the
worm pass through the feces of the green frog (Rana
clamitans) and are ingested by the pulmonate snail, Helisoma anceps. While in H. anceps sporocysts and redia
stages develop ending in the development of cercariocysts. Feeding activity of Cypridopsis sp. transfers the
cercaria body into the hemocoel of the ostracode.
Odonate naiiads eat the affected ostracodes and, in
turn, are eaten by green frogs. Griffiths and Evans
(1994) found the infestations of the peritrich Nüchterleinella corneliae on Cypria ophtalmica and Cyclocypris ovum. Infestations where more prevalent during
winter and decreased during the summer. The parasite
tended to be clustered around the bases of the appendages, particularly the genitalia and the furcae.
Bronnvall and Larsson (1994, 1995) described new
microsporidian parasites, Flabelliforma ostracodae and
Binucleospora elongata in the ostracode Candona sp.
Their investigations, based on light microscopic and ultrastructural characteristics found the musculature, the
frequent site for the development of microsporidia, was
undisturbed. The infection was restricted to the adipose
and connective tissues and to the haemocytes which appeared to be filled with spores. Diarra and Toguebaye
(1996) examined sections of ostracodes infected by
Nosema stenocypris which revealed that the microsporidium develops particulary in the muscles of
Stenocypris major which are destroyed and replaced by
the developmental stages of the parasite.
XI. COLLECTING AND REARING TECHNIQUES
A. Collecting and Preservation
Ostracodes have been collected using various kinds
of gear such as a Birge cone net (Ferguson, 1958b). Using a net has certain disadvantages when gathering
benthic organisms; many true benthic forms are not
collected because the net usually does not thoroughly
sample substrates which ostracodes live on and in. Bais
and Agrawal (1995) collected zooplankton using a
plankton net (0.25 mm); however, such an apparatus
will only collect the larger ostracode species while missing a few smaller forms and the true benthic fauna. The
use of an Ekman dredge or similar device is recommended for a quantitative sample of the benthos (Delorme, 1967). Care should be taken not to allow the
dredge to settle too far into the substrate, 2 cm being
optimal. The actual depth to which a sampling device
should go may be checked out by pushing a short core
tube into the sediments to observe the depth to which
830
L. D. Delorme
ostracodes have penetrated. Any dredge that is used
should have flaps on the top so water will flow through
the dredge when it is being lowered. This will prevent a
bow wave from preceding the dredge and disturbing
the sediment – water interface. When the dredge is retrieved, the flaps will close and prevent the interface
and collected organisms from being disturbed and
swept away by moving water.
The large quantity of sediment (500 cm3) retrieved using a dredge means that a method of concentrating the ostracodes must be used. The sample can be
washed through a set of 8-in. sediment sieves (#20
mesh, 0.841 mm; #60 mesh, 0.25 mm; #100 mesh.
0.149 mm) using a gentle stream of water from a sprinkler head. This procedure removes particle size fractions
(including instars) finer than fine sand (0.149 mm). The
smallest freshwater ostracode is Limnocythere friabilis
with a height of 0.23 – 0.31 mm. The adult shells of this
species are readily retained by both the #60 and #100
mesh. Large cyprids, such as Cypris pubera, megalocyprids, herpetocyprids, and prionocyprids will be concentrated on the #20 mesh sieve. When the specimens
are used to study appendages and other organs, ostracodes can be picked by hand from the washed residue
by using a steromicroscope. The specimens should then
be placed in 5 – 10% buffered formalin solution. Concentrated formalin, ethanol, and methanol should not
be used, as these preservatives will decalcify the shell. It
has been suggested that sodium tetraborate (250 cc saturated solution) be added to 20 L of 75% ethanol to
prevent calcitic shells from being dissolved. Specimens
may be mounted in Hoyer’s solution, glycerin, or
polyvinyl lactophenol. However, with the use of the
scanning electron microscope (SEM) to study appendages, these mounting media are not necessary.
When preservation of the visceral mass or the appendages is not critical, freeze drying is suggested to
preserve the specimens for easy storage. The residue
from the sieves is transferred to a beaker using a spatula and a small amount of water from a wash bottle.
The sediment is then frozen. After freezing, filter paper
or a suitable tissue is placed on the beaker with an elastic band and set into a freeze-drying apparatus. In the
freeze-dryer, ice is sublimed by vacuum from the specimens and sediments. This process takes 24 – 48 h, depending on the volume of ice being removed, the number of samples, and the size of the freeze-dryer. The
specimens are then handpicked from the sample using a
steromicroscope and placed in a separate vial or a micropaleontologic slide. This method has the added advantage of not requiring the more extensive labor and
time curating wet samples. The dried specimens are
also in excellent condition for further study with an
SEM to view and photograph appendages.
When a large volume of sediment still remains in
the sieves after washing and freeze drying, the ostracodes can be concentrated using acetone (Delorme,
1967). [Acetone should only be used under a safe operating fume hood.] The residue from the #60 and #100
mesh sieves is passed through a 3-in. filter sieve to ensure the sediment particles are disaggregated. The
sieved residue is then sprinkled into a funnel (4-in.)
containing acetone. The sediment particles sink and the
ostracode carapaces float to the surface. Normally, the
carapace will trap an air bubble which makes the shell
buoyant. The lower tip of the funnel is equipped with a
latex tube and a clamp. After the sediment has been
sprinkled into the acetone, the clamp is carefully removed from the tube and the “sink” allowed to drain
into another funnel lined with rapid-draining filter paper. Care must be taken not to drain all the acetone
from the top funnel, otherwise the ostracodes will be
removed. When the sink has been removed, another
funnel lined with filter paper is placed beneath, the
clamp removed, and the top funnel rinsed with acetone
from a wash bottle. In this way, all supernatant particles (the float), including the ostracodes, will be transferred to the lower filter paper. Individual shells will
also be a part of the float except Cytherissa, Darwinula, Cytheromorpha,and Limnocythere which do not
have an anterior vestibule. The filter paper containing
the ostracode residue is then air- or oven-dried, and the
residue transferred to a vial. The shells and carapaces
are picked with a very fine wetted sable hair brush and
mounted onto a micropaleontological slide. The slide is
preglued with the water-soluble gum Tragacanth. The
ostracodes are then ready to be identified and counted.
For an accurate count of adult ostracodes, the foregoing method is adequate. If, however, all live instars
and adults are required for biomass studies, then the
complete sediment parcel will be required. Wet sieving
would only assist in separating organisms and particles
by size. For biomass studies, it might be preferable to
use a short core tube pushed into the sediment some
10 – 15 cm. Nalepa and Robertson (1981) have recovered ostracodes using two mesh sizes, 0.595 and
0.106 mm, for biomass studies. The smallest mesh size
required to collect all the ostracodes including instars is
not indicated by them, as 0.106 mm will not retain all
the instars. Hummon (1981) recommends a sieve size
of 0.062 mm to retain all ostracodes for quantitative
analyses. This sieve size would miss the first three instars of Limnocythere friabilis.
B. Rearing Techniques
Cosmopolitan ostracodes, such as Cypridopsis
vidua and Cyprinotus incongruens, are relatively easy
20. Ostracoda
to rear in an aquarium. Collection of water, from the
habitat associated with the species to be reared, along
with some of the substrate will allow the population to
develop. Natural water from the habitat is preferred to
distilled or tap water. Habitat water contains the
proper mix of major and minor ions for an established
solute composition. The pH of the water should be
checked periodically and adjusted when necessary. This
type of aquarium will probably go “wild” and anoxic
after several months. A more pristine culture can be set
up by filtering the water from the habitat, placing it in
an aquarium with sterilized silt, sand, and clay as a
substrate. Cultured algae (Klugh, 1927; Kesling,
1951a; Sanders, 1970; Grant et al., 1983; Havel and
Talbot, 1995; Roca et al., 1993; Roca and Wansard,
1997; Zelmer and Esch, 1998) have been used for food
as well as fresh leaf lettuce and grated boiled egg which
has been allowed to decompose bacterially for a
month. Roca and Wansard (1997) also used benthic algae, gastropod pellets, and Spirulina with a high survival rate whereas chopped alfalfa as food with garden
compost produced the lowest survival rate. Aquaria
should not be placed on a window ledge where high
temperatures may destroy the artificial habitat.
XII. USES OF FRESHWATER OSTRACODA
Ostracode shells are preserved as fossils. Autecological data bases are available for extant species, and
so these crustaceans can be used to reconstruct past
chemical and physical habitats.
With a suitable time frame, the paleolimnology of
a pond or lake may be reconstructed from the fossils
extracted from a sediment core. Initially, subjective interpretations were made of past conditions of the lake
or pond (Gutentag and Benson, 1962; Benson and
MacDonald, 1963; Staplin, 1963a, b; Neale, 1988). As
autecological data bases were generated, paleointerpretive models were developed that used autecological
data for objective reconstruction of the chemical and
physical aspects of the habitat (Anderson et al., 1985;
Cvancara et al., 1971; Delorme, 1968, 1969b, 1971b,
c, 1982, 1996; Delorme and Zoltai, 1984; Delorme
et al., 1977, 1978; Forester et al., 1987; Karrow et al.,
1975, 1995; Klassen et al., 1967; McAllister and
Harrington, 1969; Smith et al., 1992; Westgate et al.,
1987; Westgate and Delorme, 1988; Porter et al.,
1999). In some instances, it is possible to use a number
of different fossils (ostracodes, molluscs, pollen, and diatoms) to obtain comprehensive paleoenvironmental
interpretations (Maher et al., 1998). A summary of the
use of Canadian ostracodes in paleolimnology and paleoclimatology is given by Delorme (1989).
831
Forester et al. (1994) have devised two interesting indices which estimate the “TDS ratio” of evaporation to
outflow of a large lake and a “shore – zone ratio” which
gives an indication of the proximity of the shore line to
the core site. In each case, ostracode abundances are used
in the ratios. Curry (1999) has devised two environmental tolerance indices (ETI) based on 341 modern aquatic
environments in the United States. The indices are based
on a limited set of autecological data and so at this time
cannot be universally applied for North America.
Trace element analyses of Mg, Ca, and Sr in ostracode shells have been used for reconstructing past salinity changes in a variety of aquatic habitats (Chivas
et al., 1983, 1985, 1986a, b; De Deckker, 1983b, 1988;
Lamb et al., 1995). Similar studies are being carried out
on North American ostracode shells. Engstrom and
Nelson (1991) found that distribution coefficients (KD)
for (Mg2 /Ca2)H2O and (Sr2 /Ca2)H2O in shells of
Candona rawsoni from cultured samples and from
shells collected from Devils Lake, North Dakota have
similar ratios. Analyses show excess Mg is incorporated
into the shell during early calcification which is a confirmation of the work done by Chivas et al. (1986a). Hu
et al. (1998) did trace-element analyses of ostracode
shells from a sediment core of Farewell Lake, Alaska.
From the Mg / Ca and Sr / Ca ratios they detailed the climatic reconstruction of the area indicating cold and
warm climate periods. A 19-m core from Coldwater
Lake, North Dakota documented several century-scale
changes in the paleoenvironmental record using 18O,
13C, Mg / Ca, and Sr / Ca ratios (Xia et al., 1997b). A
total of seven phases of climate and salinity were identified. Yu and Ito (1999) using Candona rawsoni shells
from a Rice Lake, North Dakota, core determined
Mg / Ca ratios showing periods of salinity and aridity.
Oxygen isotope studies of calcitic shells are being
used to develop paleoclimatic interpretations (von
Grafenstein et al., 1992, 1994). Middle Holocene
ostracodes of Elk Lake, in Grant County, Minnesota,
show an assemblage dominated by Limnocythere
staplini, a halophyte. This species is a high-salinity
form, yet the oxygen isotope values on shells of Candona rawsoni indicate a decrease by 2 – 3 ppt indicating
that ground water mediated the paleoclimatic record.
Xia et al. (1997a, c) have determined that calcitic shells
of Candona rawsoni formed at temperatures of 15 and
25°C were not in isotopic equilibrium with water, but
had a constant offset from equilibrium based on oxygen isotope fractionation of about 2 ppt. With the
molting process occurring from spring to fall, the intraannual temperature change would exceed the interannual to century-scale variations expected during lateQuaternary climate shifts. The resulting intraannual
signal may exceed the long-term paleoclimatic signals.
832
L. D. Delorme
The authors suggest 10 – 15 shells per sample be analyzed to ameliorate the interannual noise.
XIII. CURRENT AND FUTURE
RESEARCH PROBLEMS
The European literature on freshwater ostracodes
continues to grow. Unfortunately, the North American
literature is not doing as well. There is a real need for a
comprehensive review and update of North American
systematics. Detailed descriptions of soft and hard part
morphology is desperately needed together with detailed
drawings and SEM photographs. This information is
critical if scientists hope to be able to continue to communicate about the biological units on which research is
being conducted. Are we talking about the same species?
Specific research into ostracode systematics needs to
be emphasized. This is particularly true of several generic
pairs such as Cypricercus and Eucypris, Cyprinotus and
Heterocypris, and Megalocypris and Cypriconcha. These
groups require a detailed cytogenic and SEM study of
shell and appendage morphology of species of the world
(see Section XIV.A). Only in this way will we be able to
resolve which biocharacters of a certain group should be
used in defining a genus. The establishment of a universally acceptable systematic nomenclature is important for
studies in ecology and paleolimnology.
Continuing studies are required in autecology of
ostracodes. Data bases of chemical and physical parameters of ostracode habitats are being expanded (B. B.
Curry, R. M. Forester, A. J. Smith, personal communication; L. D. Delorme unpublished). Analyses of these
data bases are required to give a sense of the factors
which control the species in its environment. The roles
of solute composition and salinity of the habitat are an
integral part of this analysis.
The role of ostracodes in the population dynamics
of a pond or lake system is poorly known. Only a few
studies have attempted to look at biomass (Bechara,
1996), productivity, densities in communities, positions
in the aquatic food chain (as predators or prey), behavioral defense mechanisms, and host – parasite interactions. All these factors and others may affect the success of ostracodes in their habitat and need to be
studied in more detail. More scientific papers have been
written since the first edition of this book, but more are
needed as many more questions are being raised.
XIV. CLASSIFICATION OF OSTRACODES
Ostracodes are typically identified from adult specimens. Insufficient information is obtained from the instars (juveniles) to make a proper identification to the
species level unless a monoculture has been obtained
and adults are present. In some instances, identification
of the instar can be taken to the generic level. The most
useful biocharacter in taxonomy and systematics is the
shell. A field key is offered by Delorme (1967) for some
of the North American ostracode species as well as his
five-part series (Delorme, 1970a – d, 1971a). Other useful references are Furtos (1933) and Hoff (1942).
A. Taxonomic Controversies
A number of controversies exist around the naming of certain genera. These will be discussed separately. In order not to confuse the issue any further, a
conservative approach will be taken for the use of
generic names. Detailed systematics using the scanning
electron microscope will be required, using specimens
from the global arena rather than from local areas, in
order to resolve the issues.
The names of two genera Cypricercus and Eucypris
have been commonly used. Broodbakker (1983) has provided a literature review of the status for Cypricercus and
Strandesia. There is no mention of the genus Eucypris.
Sars (1928) differentiated between Eucypris and
Cypricercus on the absence and presence of males respectively. He also noted the peculiar arrangement of the
“spermatic vessels” (Sars, 1928, p. 117) in the males of
Cypricercus. Furtos (1933) maintained the two genera
based on reproduction and length of the furcal ramus
(1 / 2 the length of shell, Eucypris; 1 / 2 the length of
shell, Cypricercus). Hoff (1942) noted the elevation of
genera on the basis of propagation was not an acceptable
practice. He opted for one genus based on an extremely
long furcal ramus for Cypricercus. The species Eucypris
crassa was originally assigned to Cypris by O. F. Müller
and later to Eucypris by G. W. Müller (1912). This
species has always been referred to the genus Eucypris.
Martens (1989) has ascribed Eucypris serrata to the
genus Trajancypris, but has not included the North
American form (Delorme, 1970a) to the new genus.
Broodbakker (1983) erected the genus Bradleystrandesia
to include those species of Cypricercus which occur in
Europe and North America. This author states “further
research is needed to prove if the genus Bradleystrandesia
is confined to Europe and North America” (Broodbakker, 1983, p. 329). Martens (in Turgeon and Hebert,
1995) continued to use Cypricercus for the North American species. Bradleystrandesia has not been confirmed
for North America, but has been used by Rossi et al.
(1998) in referencing work done by Turgeon and Hebert
(1995). Cypricercus and Eucypris will be used in the key.
The genera Cyprinotus Brady and Heterocypris
(Claus) have had a confused nomenclature. Purper and
Würdig-Macial (1974) did a literature review of these
two genera. They concluded the main difference
20. Ostracoda
between the two genera was the dorsal gibbosity of the
right valve for Cyprinotus. Both genera are reported as
having the anteroventral margin of the right valve with
denticles or crenulations. Sars (1928) and Müller
(1912) did not consider the gibbosity of the right valve
as a valid generic biocharacter and, thus, did not recognize Heterocypris. Hoff (1942) also did not recognize
Heterocypris. This morphological biocharacter in
Physocypria (Delorme, 1970b) may or may not be present and is not used to split the genus. Cyprinotus is
used in the key because it was published first in 1886.
The genus Cypriconcha was erected by Sars (1926)
to incorporate the large cyprinids of North America.
Sars (1898) had already established Megalocypris for
the large cyprinids of Africa. Delorme (1969a) reviewed the two genera and concluded the similarities
were many and discrepancies few. As a result he put
the North American cypriconchids into the genus
Megalocypris. Martens (1986) and McKenzie (1982,
referenced in Martens, 1986) argued for the retention
of Cypriconcha for the large cyprinids of North America. Megalocypris is used in the key.
A number of genera may be found in the North
American ostracode literature, but are not included
here. The following generic names are no longer considered valid:
Cyclocypria Dobbin, 1941 — Cyclocypris Brady and
Norman, 1888
Cypriconcha Sars, 1926 — Megalocypris Sars, 1898
[see Delorme, 1969a]
Cytherites Sars, 1926 — Entocythere Marshall, 1903
[see Hoff, 1943b]
Pionocypris Brady and Norman, 1896 — Cypridopsis Brady, 1868
Pseudoilyocypris Ferguson, 1967 — Pelocypris Klie,
1939 [see Delorme, 1970d]
Spirocypris Sharpe, 1903 — Cypricercus Sars, 1895
[see Hoff, 1942]
833
Several species, as listed below, have been assigned
to an incorrect genus; therefore, the generic names will
not appear in the key:
Candonocypris deeveyi Tressler, 1954 — Megalocypris deeveyi (Tressler) 1954
Candonocypris pugionis Furtos, 1936b — Megalocypris[?] pugionis (Furtos), 1936b [see Furtos, 1936b]
Candonocypris sarsi Danforth, 1948 — Megalocypris
sarsi (Danforth), 1948
Candonocypris serrato-marginata (Furtos), 1935 —
Eucypris serrato-marginata (Furtos), 1935
Erpetocypris barbatus Turner, 1899 — Megalocypris
sp. (Turner), 1899
Prionocypris canadensis Sars, 1926 — Strandesia
canadensis (Sars), 1926 [see Marmonier and Ward,
1990]
Prionocypris glacialis (Sars), 1890 — Tonnacypris
glacialis (Sars), [see Griffiths et al., 1998]
Pseudocandona ikpikpukensis Swain, 1963 —
Candona ikpikpukensis (Swain), 1963 [see Delorme,
1970c]
Stenocypria longicomosa Furtos, 1933 — Isocypris
longicomosa (Furtos), 1933 [see Delorme, 1970a]
B. Classification of the Order Podocopida
The Podocopida1 is the only order that contains
both marine and freshwater ostracodes (Table II). Dorsal border of the valve is curved, or if straight, it is
much shorter than the total length. No permanent anterior opening is present between the valves. There is
no heart; two or three pairs of thoracic legs are present.
Within Podocopida are two suborders: Metacopina
and Podocopina. As will be described below, the suborder Metacopina contains only one freshwater genus in
North America, Darwinula (superfamily Darwinuloidea, family Darwinulidae; Figs. 8 and 10). Its characteristics are given as follows2.
Shell surface smooth, vestibules absent, distinctive adductor muscle scars (Figs. 8, 10, 19); first antenna composed of six podomeres with strong
spikelike setae, swimming setae lacking on second antenna, mandibular exopodite with short palp of three podomeres of which the first is wide
and has a row of long feathered setae, respiratory plate of mandible small, masticatory process of maxilla short and heavy, respiratory plate of
maxilla with numerous feathered setae, maxillae with strong masticatory structure and leglike palp of three podomeres, first and second thoracic legs similar in structure and direction, furca lacking, ovaries do not originate between inner and outer epidermis, female carries eggs in
posterior of carapace ..............................................................................................................................................................................Darwinula
Freshwater and some marine ostracodes are
included in the suborder Podocopina. Members of this
taxon have two pairs of thoracic legs. The exopodite of
the second antenna is represented by at most a scale
and a few setae. The furcae are rodlike rather than
lamelliform; eyes are present in most forms. Closing
muscle scars are discrete and arranged in a distinctive
group. Taxonomic keys to genera within two superfamilies of this suborder are given below.
C. Taxonomic Key to Genera of the
Superfamily Cypridoidea2
Surface of the shell usually smooth, dorsal margin
without interlocking teeth. Eyes developed to varying
degrees, either separated or fused into a single median
eye. The first antennae with basal portion of two or
1
2
Modified after Kesling (1951a).
Modified after Hoff (1942).
834
L. D. Delorme
TABLE II
Classification of the Class Ostracodaa
Class Ostracoda
Subclass Podocopa
Order Podocopida Sars, 1866
Suborder Metacopina Sylvester-Bradley, 1961
Superfamily Darwinuloidea Brady and Norman, 1888
Family Darwinulidae Brady and Norman, 1888
Genus Darwinula Brady and Norman
Suborder Podocopina Sars, 1866
Superfamily Cypridoidea Baird, 1845
Family Candoniidae Kaufmann, 1900
Subfamily Candoninae Daday, 1900
Genus Candocyprinotus Delorme
Genus Candona Baird
Genus Fabaeformiscandona Krstic
Genus Nannocandona, Eckman
Genus Paracandona, Hartwig
Genus Pseudocandona, Kaufmann
Family Cyprididae Baird, 1845
Subfamily Cypridinae Baird, 1845
Genus Chlamydotheca Sausseure
Genus Cyprinotus Brady
Genus Cypris Müller
Genus Dolerocypris Kaufmann
Genus Herpetocypris Brady and Norman
Genus Ilyodromus Sars
Genus Isocypris Müller
Genus Megalocypris Sars
Genus Scottia Brady and Norman
Genus Stenocypris Sars
Genus Strandesia Stuhlmann
Subfamily Eucypridinae Bronstein, 1947
Genus Cypricercus Sars
Genus Eucypris Vávra
Genus Tonnacypris Diebel and Pietrzeniuk
Genus Trajancypris Martens
Subfamily Cyclocyprinae Hoff, 1942
Genus Candocypria Furtos
Genus Cyclocypris Brady and Norman
Genus Cypria Zenker
Genus Physocypria Vávra
Subfamily Cypridopsinae Kaufmann, 1900
Genus Cavernocypris Hartmann
Genus Cypretta Vávra
Genus Cypridopsis Brady
Genus Potamocypris Brady
Genus Sarscypridopsis McKenzie
Family Ilyocyprididae Kaufmann, 1900
Subfamily Ilyocyprinae Kaufmann, 1900
Genus Ilyocypris Brady and Norman
Genus Pelocypris Klie
Family Notodromadidae Kaufmann, 1900
Subfamily Notodromadinae Kaufmann, 1900
Genus Cyprois Zenker
Genus Notodromas Lilljeborg
Superfamily Cytheroidea Baird, 1850
Family Cytheridae Baird, 1850
Subfamily Cytherideinae Sars, 1928
Genus Cyprideis Jones
Family Entocytheridae Hoff, 1942
Subfamily Entocytherinae Hoff, 1942
Genus Ankylocythere Hart
(Continues)
TABLE II
(Continued)
Genus Ascetocythere Hart
Genus Cymocythere Hart
Genus Dactylocythere Hart
Genus Donnaldsoncythere Hart
Genus Entocythere Marshall
Genus Geocythere Hart
Genus Harpagocythere Hobbs III
Genus Hartocythere Hobbs III
Genus Litocythere Hobbs and Walton
Genus Lordocythere Hobbs and Hobbs
Genus Okriocythere Hart
Genus Ornithocythere Hobbs
Genus Phymocythere Hobbs and Hart
Genus Plectocythere Hobbs III
Genus Rhadinocythere Hart
Genus Saurocythere Hobbs III
Genus Sagittocythere Hart
Genus Thermastrocythere Hobbs and Walton
Genus Uncinocythere Hart
Family Limnocytheridae Klie, 1938
Subfamily Limnocytherinae Klie, 1938
Genus Limnocythere Brady
Genus Metacypris Brady and Robertson
Family Loxoconchidae Sars, 1928
Subfamily Loxoconchinae Sars, 1928
Genus Cytheromorpha Hirschmann
Family Neocytherideidae Puri, 1957
Subfamily Neocytherideidinae Puri, 1957
Genus Cytherissa Sars
a
The systematic list above is alphabetical (after Bowman
and Abele, 1982), but the systematic treatment is not.
three podomeres and an endopodite of four or five
podomeres, with swimming setae usually well developed. The second antennae with basal part of two
podomeres and an endopodite of three or four
podomeres. The exopodite is reduced to a small scalelike appendage bearing at most three setae. Maxillula
not pediform, but modified as a mouth part, with the
anterior margin of the base adapted for feeding. The
maxillar endopodite forms a small palp in the female,
but is enlarged to form prehensile palps in the male.
The first thoracic leg has an endopodite of three or four
podomeres and a strong distal claw. The second leg is
bent dorsally and is probably used in cleaning the
respiratory surfaces and other parts of the body. The
second leg usually has three distal setae, but the distal
end may be modified for grasping. The furca is typically well developed and rod-shaped, but may be reduced to a flagellum or whiplike structure. The gonads
are located within the valves of the shell. In the male, a
portion of the vas deferens is modified to form an ejaculatory duct.
FIGURE 19 External view of Darwinula stevensoni (Brady and Robertson). FIGURE 20 External view of
Cypridopsis vidua (Müller). FIGURE 21 External view of Potamocypris smaragdina (Vávra). FIGURE 22
External view of Cyprois marginata Straus. FIGURE 23 External view of a female Candona rawsoni Tressler,
note the overlap of the right valve by the left valve. FIGURE 24 External view of Paracandona euplectella
Brady and Norman, note the reticulate structure and the pustules. FIGURE 25 External view of Cypricercus
reticulatus (Zaddach).
835
836
L. D. Delorme
1a.
Furcal ramus greatly reduced, whip-shaped, without terminal claw (difficult to observe)Cypridopsinae3 ...........................................2
1b.
Furcal ramus well developed, bar-shaped, with two terminal or subterminal claws; outer masticatory process of maxilla with six
nearly equal setae modified to form toothed spines ..............................................................................................Notodromadinae4 4
1c.
Swimming setae of antennae completely lacking, shell white; outer masticatory process with two or three setae modified as spines
.....................................................................................................................................................................................Candoninae5 5
1d.
Swimming setae of first antenna present; second thoracic leg distally modified as seizing apparatus, ultimate podomere beak-like
with two well developed bristle-like setae, third setae lacking or hook-like....................................................................Cypridinae6 7
1e.
Third thoracic leg with one long reflexed seta and two shorter unreflexed setae .....................................................Cyclocyprinae7 10
1f.
Carapace 1. 0 – 2.5 mm, elliptical to elongated, rarely globular; surface ornamentation absent except for shallow pits; marginal
teeth possible on both valves; marginal zone with selvage present or absent, sometimes inwardly displaced in the right valve. First
antenna with Rome organ small and undivided; mandibular palp with gamma-seta short, stout and hirsute. Maxillula with third endopodite carrying two, mostly smooth, teeth or bristles. The second process of the maxillula with “c”-seta ............Eucypridinae8 12
1g.
Second thoracic leg not bearing chela, last podomere cylindrical and bearing three setae; shell oblong to subrectangular, dorsal
margin straight, surface pitted, with one or more transverse sulci ................................................................................Ilyocyprinae9 4
2a(1a).
Shells tumid........................................................................................................................................................................................3
2b.
Shells crescentic in shape, compressed, right valve higher than left and extending dorsally above left, H / L 0.5, right valve overlaps
left valve in venter, usually hairy; swimming setae of second antenna well developed, usually extending to tips of claws or beyond;
ultimate podomere of maxillary palp distally wider than long (Fig. 21) ..........................................................................Potamocypris
3a(2a).
Shells subovate, H / L 0.5, valves nearly equal, left valve overlaps right valve in venter; ultimate podomere of maxillary palp cylindrical, longer than wide (Fig. 20), swimming setae of second antennae well developed......................................................Cypridopsis
3b.
Shells subtriangular, H / L 0.5, right valve overlaps left valve in venter; swimming setae of second antennae well developed
.....................................................................................................................................................................................Sarscypridopsis
3c.
Shells elongate, 1 mm in length, left valve longer than right valve, H / L 0.5, left valve overlaps right valve in venter; swimming
setae of second antenna poorly developed, furcal rami absent in males..........................................................................Cavernocypris
3d.
Shells tumid, 1 mm in length, anterior margin of both valves with row of radiating septa; two maxillary spines may be smooth or
toothed; ovaries or testes coiled in spiral pattern in posterior of carapace...............................................................................Cypretta
4a(1b).
Shell from side short and high, compressed; eyes not widely separated; first antenna of five podomeres; swimming setae almost
reach tips of end claws; right and left prehensile palps of male maxillar endopodites differing little, with well-developed respiratory
plate; setae on distal end of second thoracic leg modified for grasping; furca with two claws and two setae (Fig. 22) ..............Cyprois
4b.
Shell short with height two-thirds of length, dorsal margin forming elevated arch, ventral surface flattened; eyes well separated; first
antenna of six apparent podomeres, swimming setae extending beyond tips of terminal claws; palps of maxillae in males not similar,
each formed with two podomeres; maxillae without respiratory plate; distal three setae of second thoracic leg nearly equal; furca
with terminal seta lacking.................................................................................................................................................Notodromas
5a(1c).
Carapace with sides convex; outer masticatory process of maxilla smooth; apical claw of ultimate segment of second thoracic leg
well developed, furcal claws weakly serrated .............................................................................................................Candocyprinotus
5b.
Shell subrectangular to subtriangular in shape, two special sensory setae present at juncture of fourth and fifth podomeres of male
first antenna, respiratory plate of maxillae with two or three setae ....................................................................................................6
6a(5b).
Male carapace shape and size usually different than female; second thoracic leg with three unequally long setae on last podomere;
Zenker’s organs usually with seven wreaths of chitinous spines, openings of ejaculatory duct funnel-shaped (Figs. 9a, 9b, 23)
..............................................................................................................................................................................................Candona
6b.
Carapace usually elongate, more rarely triangular in lateral view,distinctly compressed in dorsal view; left valve usually with a
posterodorsal lobelike extension of the shell which overlaps right valve; male second antenna dimorphic, penultimate segment with
male bristle. Setal group of the 2nd segment of the mandibular palp with three (fabaeformis group) or four (acuminata group) setae,
gamma seta of the penultimate segment smooth (not plumose). Protopodite of second thoracic leg with two setae (d1 and dp), setae
‘e’ and ‘f’ of first endopodite missing, seta ‘g’ of third endopodite present, terminal endopodite with two long (h2 and h3) and one
3
Modified after Marmonier et al. (1989), and McKenzie (1977).
Modified after Hoff (1942).
5
Modified after Hoff, 1942, Furtos (1933) and Meisch (1996).
6
Modified after Hoff (1942) and Delorme (1970a).
7
Modified after Hoff (1942) and Furtos (1933).
8
Modified after Martens (1989).
9
Modified after Hoff (1942) and Tressler (1949).
4
20. Ostracoda
837
short (h1) setae. Female genital lobe usually with more or less well-developed process directed posteriorly. Hemipenis with ‘M’
process heavily sclerotized, proximal part slightly broader than the distal one.....................................................Fabaeformiscandona
6c.
Shell usually with small tubercles and reticulations on surface, shape subrectangular; penultimate segment of second thoracic leg divided, each division with strong distal seta; otherwise similar to Candona (Figs. 7, 24) ...................................................Paracandona
6d.
Carapace of medium length (0.8 – 1.2 mm), variously shaped, usually short and stout, more rarely elongate (subrectangular) or triangular in lateral view. Surface of adult valves smooth or pitted, usually with long stiff setae emanating through the shell, left valve
overlaps the right valve ventrally. The penultimate endopodite of the second antennae subdivided or not, when subdivided with
male bristles (t2 and t3) transformed. Setal group of the second endopodite of mandibular palp with three or five setae, distal seta
of penultimate endopodite smooth (not plumose). Protopodite of second thoracic leg with three setae (d1-d2, dp), distal seta ‘g’ on
the penultimate endopodite present, terminal endopodite with one short (h1), and two long setae (h2-3). Zenker’s organ with 5 2
rings of spines. Hemipenis with ‘M’ process flat and proximally only weakly sclerotized ............................................Pseudocandona
7a(1d).
Margins of valve usually smooth ........................................................................................................................................................8
7b.
Margin of valve with spines or denticles...........................................................................................................................................19
8a(7a).
Shells 2.2 mm long ..........................................................................................................................................................................9
8b.
Shells 2.2 mm long ........................................................................................................................................................................16
9a(8a).
Shell surface sculptured ....................................................................................................................................................................10
9b.
Shell surface smooth.........................................................................................................................................................................13
10a(9a).
Shell surface reticulate or scabrous especially in juveniles.................................................................................................................11
10b.
Shell surface punctate or striated ......................................................................................................................................................12
11a(10a).
Outer masticatory process with two long, toothed spinelike setae; furcal ramus greater than half length of valve; testes coiled in anterior portion of each valve, males rare (Fig. 25)................................................................................................................Cypricercus
12a.
Outer masticatory process with two long, toothed spinelike setae; furcal ramus less than half length of valve, males rare (Fig. 26)
...............................................................................................................................................................................................Eucypris
12b.
Swimming setae of second antenna rudimentary, length of gamma-seta of the mandibular palp is nearly five times its basal width,
second segment of mandibular palp is trapezoidal and the terminal endopodite of the third endopodite bears two serrated teeth, first
thoracic leg has the setae d2 nearly two times d1; shell elongate and moderately large, compressed and subrectangular to clavate, anterior inner lamellae of left valve with small peg tooth sometimes nearly invisible; furcal ramus smooth ..........................Tonnacypris
12c.
Right valve with a frontal selvage inwardly displaced to a varying degree, but never marginal; left valve without selvage, but with
large frontal inner list, situated about halfway on the calcified inner lamella ....................................................................Trajancypris
13a(10b).
Shell elongate, narrow with scattered punctae; swimming seta of second antenna barely reaching tips of claws, second thoracic leg
with well-developed apical claw; rami of furcae dissimilar in width, two end claws strongly pectinate...............................Stenocypris
13b.
Shells elongate and compressed, surface longitudinally striated; swimming setae of first and second antenna short, furcal ramus terminates in three short claws rather than two ......................................................................................................................Ilyodromus
14.
Swimming setae of second antenna strongly developed ....................................................................................................................15
15a(14b).
Claws of furca finely denticulate; shell subrectangular, surface smooth and moderately hairy; characteristic marginal pore canals in
anterior duplicature; outer masticatory process of maxilla with two smooth spines; second thoracic leg with well-developed apical
claw .......................................................................................................................................................................................Isocypris
15b.
Claws of furca strongly developed....................................................................................................................................................16
16a(15b).
Shell reniform, densely hairy; maxillae bears well-developed respiratory plate; first thoracic leg with one long terminal seta and
claw, furca with feathered dorsal seta ........................................................................................................................................Scottia
16b.
Shells elongate, dorsum often with prominent dorsal flange, left valve always with conspicuous row of tuberclelike canals removed
from the free margin; swimming setae of second antenna extend to tips of terminal claws, third masticatory process of maxilla with
two strong spines, furcal ramus at least half as long as valve and with two claws and setae .................................................Strandesia
17a(8b).
Shells with anterior flangelike projections which produces large vestibule; both antenna with well-developed swimming setae; outer
masticatory process of maxilla with one toothed and one denticulate spine; second thoracic leg with well-developed apical claw;
claws of furca finely denticulate ...................................................................................................................................Chlamydotheca
17b.
Shells without anterior flangelike projection.....................................................................................................................................18
18a(17b).
Shells narrow and spindle-shaped, vestibules well developed at both extremities; both antennae with well-developed swimming setae; maxilla with outer masticatory process with two smooth spines; second thoracic leg with well-developed apical claw, claws of
furca coarsely denticulate .................................................................................................................................................Dolerocypris
838
L. D. Delorme
FIGURE 26
External view of Eucypris crassa (Müller). FIGURE 27 External view of Cypris pubera Müller,
note the posteroventral spines on the right valve. FIGURE 28 External view of Cyclocypris ampla Furtos,
note the pitted surface. FIGURE 29 Internal view of Physocypria pustulosa (Sharpe) showing the denticles on
the anteroventral margin. FIGURE 30 Exterior view of Pelocypris alatabulbosa Delorme showing a reticulate
surface, sulci, and nodes.
18b.
Shells subrectangular ........................................................................................................................................................................19
19a(18b).
Shells elongate and large, compressed; swimming setae of second antenna not reaching mid position of terminal claw; spines of
outer masticatory process of maxilla toothed; ventral edge of furca denticulate with combs of teeth .............................Herpetocypris
19b.
Shells subrectangular to subtrapezoidal; swimming setae on first antenna extend to end of ultimate podomere and not to tips of
claws of second antenna; outer masticatory process made up of short, smooth bristles; maxillar endopodite of male developed into
prehensile palp; second thoracic leg with apical claw not well developed; claws of furca denticulate...............................Megalocypris
20. Ostracoda
839
20a(7b).
Shells 2 mm, margin of right or left valve tuberculate and margin of other valve smooth, dorsal margin may be arched or have an
obtuse apex, valves commonly unequal, anterolateral surfaces pustulose; outer masticatory process of maxilla with two clawlike setae which may or may not be toothed; furcal claw longer than one-half length of ventral margin of furcal ramus, ramus with length
less than twenty times least width (Figs. 3, 4) .....................................................................................................................Cyprinotus
20b.
Shells 2 mm, tumid, highly arched, venter flattened, anterior margins denticulate, posteroventral margin of right valve with two to
four sharp marginal spines, surface wrinkled to minutely crenulate to punctate; second antenna with well-developed swimming setae; outermost masticatory process of maxilla with two denticulate spines, second thoracic leg with well-developed apical claw,
claws of furca strongly denticulate (Fig. 27) ...............................................................................................................................Cypris
21a(1e).
Swimming setae of second antenna lacking or rudimentary; shells ovoid, compressed; eyes well developed and fused; ultimate segment of second thoracic leg with two short and one long reflexed setae; tips of furcal claws serrate ................................Candocypria
21b.
Swimming setae of second antenna well developed ..........................................................................................................................21
22a(21b).
Shells tumid, height and width greater than one-half length, surface smooth and usually brown in color; eye well developed; respiratory plate of maxillae with six plumose setae; second thoracic leg consisting of four podomeres, ultimate podomere elongated and
more than twice as wide and usually at least one-half as long as penultimate podomere; ultimate podomere of third thoracic leg
with three distal setae, unequal, with outermost one very long and reflexed (Fig. 28) ........................................................Cyclocypris
22b.
Shells compressed .............................................................................................................................................................................23
23a(22b).
Shells short and high, occasionally elongate reniform, margins smooth not tuberculate; eyes well developed; second antenna of male
with penultimate podomere divided and bearing specialized male setae; ultimate podomere of mandibular palp elongated three
times as long as proximal width; palp of maxilla well developed, masticatory process weak; penultimate podomere of second thoracic leg undivided, ultimate podomere short scarcely longer than wide, longest distal seta reflexed; terminal and subterminal claws
of furca strong, dorsal seta may be rudimentary; Zenker’s organ with seven whorls of chitinous rays, proximal end much inflated;
hemipene with two terminal lobes only, outer lobe lacking........................................................................................................Cypria
23a(22b). Shells commonly unequal in height or length or both, at least anterior margin of one of the valves tuberculate; otherwise as in genus
Cypria (Fig. 29) .................................................................................................................................................................Physocypria
24a(1g).
Shells may have large rounded humplike projections on lateral surface, marginal spines and pustules; first antenna with some swimming setae shortened and clawlike; swimming setae of second antenna may be shortened, male without special setae; maxillar endopodite of male transformed into prehensile palp, in female clearly leglike; ultimate podomere in second thoracic leg cylindrical
with three setae, longest may be reflexed, claws well developed; Zenker’s organs with numerous chitinous rods and with spherical
inflated openings at each end; vasa deferentia twisted ...........................................................................................................Ilyocypris
24b.
Shells with pronounced mamilliarylike projections on lateral surface, marginal spines and pustules; second antenna with swimming
setae which extend considerably beyond tips of terminal claws; maxillar endopodite with long curved terminal claw; first leg with
one short and one long seta; furca well developed with dorsal seta longer than terminal claw (Fig. 30) ...............................Pelocypris
D. Taxonomic Key to Genera
of the Superfamily Cytheroidea10
Shell variable in shape and sculpturing, seldom
smooth, usually with reticulations, often with spines,
furrows, or tubercles. Shells noncalcareous in the Entocytherinae. Valves nearly equal; often toothlike projections along the hinge. The first antennae consist of a
base of two podomeres and an endopodite of three or
four podomeres. The setae of the first antennae are
short and stout, often clawlike. The exopodite or the
flagellum of the second antennae is represented by a
long hollow seta forming a duct carrying secretions
from a gland. The endopodite of the second antennae
1a.
10
11
of three podomeres, the long penultimate one may be
divided. Swimming setae lacking. Maxillar endopodites
and two pairs of thoracic legs similar and all adapted
for crawling except in the Entocytherinae modified for
grasping. Furca always greatly reduced. In the male,
the hemipenes always present and well developed, the
ejaculation duct is absent, a male accessory sense organ
consisting of numerous setae on a short base is located
between and somewhat medially to the bases of the
first and second pairs of thoracic legs. The gonads do
not lie between the inner and outer lamellae of the
valves, but are in the body lateral to the intestine. In
some species, the eggs are retained in the shell cavity
during development.
Free margins of valves without conspicuous pore canals, carapaces non calcareous, reniform or subelliptical in shape, furrows or
protuberances may be present; respiratory plate of mandible reduced to two or three setae, masticatory lobes well developed, furca
rudimentary, penis strongly curved; North American ostracodes commensal on crayfishes and a freshwater crab (Fig. 12)
..............................................................................................................................................................Subfamily Entocytherinae11 3
Modified after Hoff (1942) and Sars (1928).
Modified after Hoff (1942) and Hart and Hart (1974).
840
L. D. Delorme
1b.
Free margins of valves flattened, with many long marginal pore canals, subrectangular, often with protuberances or furrows,
strongly reticulate; respiratory plate of mandible well developed, furca usually with two short setae .................................. Subfamily
Limnocytherinae12 ........................................................................................................................................................................... 22
1c.
Shell variable in shape and sculpturing, seldom smooth, usually with reticulations, spines, furrows, or tubercles, valves nearly equal,
often with toothlike projections along hinge, vestibules absent; eyes distinctly separated, antennae well developed, exopodite of second antenna in form of long hollow seta carrying secretions from a gland near base of second antenna, setae of first antenna short
and stout, often claw like, swimming setae on second antenna lacking, first thoracic leg of male prehensile, three pairs of thoracic
leg similar, furca greatly reduced, hemipenes well developed, Zenker’s organs absent, testes lying in body lateral to intestine, in some
species eggs retained in body cavity during development ............................................................................ Subfamily Cytherideinae13
[Shells distinctly sculptured, sometimes with lateral protuberances, right valve with spine in posteroventral corner]............ Cyprideis
1d.
Thoracic legs successively increasing in length................................................................................................................................... 2
2a(1d).
Shell short and stout, hinge well developed; mandibular palp with terminal joint very small, respiratory plate well developed; furcal
ramus edged at tip with two bristles ..........................................................................................................Subfamily Loxoconchinae13
[Shell short and stout, surface pitted, marginal zones thickened, duplicature narrow; eyes distinctly separated; ultimate segment of
first antenna articulated and with four robust spines; second antenna with two claw like spines inside penultimate joint; masticatory
lobes of maxilla short] .................................................................................................................................................Cytheromorpha
2b.
Shells plump, surface generally sculptured; hinge well defined, distinct closing teeth lacking in anterior and posterior of hinge
line; mandibular palp slender, respiratory plate not well developed; furcal ramus extremely small, not well defined
............................................................................................................................................................................Neocytherideidinae12
[Shell club-shaped, surface rough and pitted; valves unequal with marginal zone thickened, duplicature narrow, vestibule absent,
poorly developed teeth present on hinge; eyes well defined; antennae well developed, first antenna armed in front with three
clawlike spines; respiratory plate of mandibular palp moderately well developed; maxilla with masticatory lobes short and stout;
thoracic legs robustly developed; furcal ramus forming two oval-thickened pieces placed vertically, each provided behind with two
very small bristles (Fig. 31)] .................................................................................................................................................Cytherissa
3a(1a).
Penis with prostatic and spermatic elements widely separated along much of their lengths ................................................................4
3b.
Penis simple; or if two elements recognizable, contiguous along their entire lengths...........................................................................6
4a(3a).
Ventral portion of peniferum tapering with tip of penis reaching, or almost reaching apex .............................................Plectocythere
4b.
Ventral portion of peniferum usually rounded or with one or more prominences, seldom tapering; if tapering, tip of penis never approaching apex...................................................................................................................................................................................5
5a(4b).
Ventral portion of peniferum rounded, without prominences.........................................................................................Phymocythere
5b.
Ventral portion of peniferum with one or more prominences ventrally and / or anteriorly...............................................Ascetocythere
6a(3b).
Penis directed posteroventrally from base........................................................................................................................Lordocythere
6b.
Penis directed anteroventrally from base ............................................................................................................................................7
7a(6b).
Finger guard absent............................................................................................................................................................................8
7b.
Finger guard present.........................................................................................................................................................................18
8a(7a).
Anteroventral portion of peniferum with acute beaklike projection..............................................................................Ornithocythere
8b.
Anteroventral portion of peniferum never with beaklike projection ...................................................................................................9
9a(8b).
External border of horizontal ramus of clasping apparatus with one or more excrescence ...............................................................10
9b.
External border of horizontal ramus of clasping apparatus entire or with few shallow subapical grooves........................................14
10a(9a).
Anteroventral portion of peniferum produced ventrally in rounded lobe..........................................................................................11
10b.
Anteroventral portion of peniferum never produced ventrally in rounded lobe; or if produced, apex acute or truncate ...................12
11a(10a).
Spermatic loop horizontal, peniferum distal to dorsal margin of spermatic loop at least twice as long as portion dorsal to loop,
clasping apparatus with external border bearing single tubercle and terminating in fanlike cluster of serrations..............Saurocythere
11b.
Spermatic loop vertical, peniferum distal to dorsal margin of spermatic loop much less than twice as long as portion dorsal to loop,
clasping apparatus with external border broadly serrate and terminating in annulations ................................................Okriocythere
12a(10b).
Anteroventral portion of peniferum never with conspicuous anterodorsally directed projection ....................................Ankylocythere
12
13
Modified after Hoff (1942).
Modified after Sars (1928).
20. Ostracoda
841
12b.
Anteroventral portion of peniferum with conspicuous anterodorsally directed projection ................................................................13
13a(12b).
Anteroventral portion of peniferum entire ..........................................................................................................................Geocythere
13b.
Anteroventral portion of peniferum bifid .........................................................................................................................Hartocythere
14a(9b).
Internal border of clasping apparatus with more than three teeth, apical cluster with more than two denticles, vertical ramus
straight .............................................................................................................................................................................................15
14b.
Internal border of clasping apparatus usually with no more than three teeth, if more than three, with only two apical denticles or
vertical ramus strongly convex posteriorly .......................................................................................................................................16
15a(14a).
Ventral portion of peniferum tapering to slender tip ....................................................................................................Rhadinocythere
15b.
Ventral portion of peniferum never slender nor tapering ...................................................................................................Entocythere
16a(14b).
Clasping apparatus not clearly divisible into vertical and horizontal rami, extremities directed at angle of at least 100 degrees
..............................................................................................................................................................................Donnaldsoncythere
16b.
Clasping apparatus usually divisible into vertical and horizontal rami, extremities directed at angle of no more than 90 degrees ....17
17a(16b).
Penis large, S-shaped or sinuous and with curved posteroventral thickening of peniferum giving forcipate appearance to ventral portion of peniferum ....................................................................................................................................................Thermastrocythere
17b.
Penis of moderate size, L-shaped, and never so disposed as to contribute forcipate appearance to ventral portion of peniferum
......................................................................................................................................................................................Uncinocythere
18a(7b).
Peniferum with accessory groove (except in Dactylocythere leptophylax where finger guard always slender and trifid) ...................19
18b.
Peniferum without accessory groove; finger guard never slender and trifid .......................................................................................20
19a(18a).
Posteroventral portion of peniferum terminating in barbed point...................................................................................Sagittocythere
19b.
Posteroventral portion of peniferum variable, but never ending in barbed point...........................................................Dactylocythere
20a(18b).
Ventral portion of peniferum bulbiform, clasping apparatus never extending so far ventrally as does peniferum.............Cymocythere
20b.
Ventral portion of peniferum slender or strongly flattened; clasping apparatus extending ventrally to or beyond ventral extremity of
peniferum .........................................................................................................................................................................................21
FIGURE 31
Exterior view of Cytherissa lacustris (Sars) showing
coarse reticulate surface and nodes. FIGURE 32 Exterior view of
Limnocythere itasca Cole showing reticulate surface, sulci, and alae.
842
L. D. Delorme
21a(20b).
Ventral portion of peniferum slender, terminating in small recurved projection...........................................................Harpagocythere
21b.
Ventral portion of peniferum flattened and with concave border ........................................................................................Litocythere
22a(1b).
Shells subrectangular, reticulate, with sulci, alae or pustules on surface, margins may be denticulate, no vestibules developed; males
longer, more inflated in posterior region than females; second antenna with three apical claws; respiratory plate of mandibular palp
well developed; masticatory lobes of maxilla short; thoracic legs moderately slender; furcal ramus well defined, conical in shape
with one terminal and one lateral seta; hemipene with basal part very large and protuberant in front (Figs. 11, 32 .......Limnocythere
22b.
Shells short and broad, surface pitted, right valve with hinge line toothed; first antenna five or six segmented, second antenna four
segmented, maxilla with three masticatory processes each longer than palp........................................................................Metacypris
LITERATURE CITED
Alm, G. 1915. Monographie der Schwedischen SüsswasserOstracoden nebst systematischen Besprechungen der Tribus Podocopa.
Zoologiska Bidrag från Uppsala 4:1 – 247.
Angell, R. W., Hancock, J. S. 1989. Response of eggs of Heterocypris
incongruens (Ostracoda) to experimental stress. Journal of
Crustacean Biology 9:381 – 386.
American Public Health Association Inc. (APHA). 1965. Standard
methods for the examination of water and wastewater, including bottom sediments and sludges. American Public Health
Association, Inc., New York. 769p.
Anderson, T. W., Mott, R. J., Delorme, L. D. 1985. Evidence for
a pre-Champlain Sea glacial lake phase in Ottawa Valley,
Ontario, and its implications, in: Current Research. Part A.
Geological Survey of Canada, Paper 85-1A, pp. 239 – 245.
Bais, V. S., Agrawal, N. C. 1995. Comparative study of the zooplanktonic spectrum in the Sagar Lake and Military Engineering
Lake. Journal of Environmental Biology 16(1):27 – 32.
Baltanás, A. 1992. On the use of some methods for the estimation of
species richness. Oikos 65(3):484 – 492.
Barclay, M. H. 1966. An ecological study of a temporary pond near
Auckland, New Zealand. Australian Journal of Marine and
Freshwater Research 17:239 – 258.
Bataille, K. J., Baldassarre, G. A. 1993. Distribution and abundance
of aquatic macroinvertebrates following drought in three prairie
pothole wetlands. Wetlands 13(4):260 – 269.
Batzer, D. P., McGee, M., Resh, V. H., Smith, R. R. 1993. Characteristics of invertebrates consumed by mallards and prey response
to wetland flooding schedules. Wetlands 13(1):41 – 49.
Bechara, J. A. 1996. The relative importance of water quality, sediment composition and floating vegetation in explaining the
macrobenthic community structure of floodplain lakes (Parana
River, Argentina). Hydrobiologia 333(2):95 – 109.
Bell, G. 1982. The masterpiece of nature, the evolution and genetics
of sexuality. Univ. of California Press, Berkeley, 600 p.
Benson, R. H. 1959. Ecology of recent ostracodes of the Todos Santos Bay Region, Baja California, Mexico. University of Kansas
Paleontological Contributions, Article 1:1 – 80.
Benson, R. H. 1961. Ecology of ostracode assemblages, in: Moore,
R. D., Ed., Treatise on invertebrate paleontology. Part
Q:Arthropoda 3, Crustacea, Ostracoda. Geological Society of
America. Univ. of Kansas Press, Lawrence, pp. Q56 – Q63.
Benson, R. H. 1967. Muscle-scar patterns of Pleistocene (Kansan) ostracodes; Essays in Paleontology and Stratigraphy. Raymond C.
Moore Commemorative Volume. University Kansas Department
of Geology Special Publication 2:211 – 241.
Benson, R. H. 1974. The role of ornamentation in the design and
function of the ostracode carapace. Geoscience and Man
6:47 – 57.
Benson, R. H. 1981. Form, function, and architecture of ostracode
shells. Annual Review of Earth and Planetary Science 9:59 – 80.
Benson, R. H., MacDonald, H. C. 1963. Postglacial (Holocene)
ostracodes from Lake Erie. University of Kansas Paleontological
Contributions 4:1 – 26.
Benzie, J. A. H. 1989. The distribution and habitat preference of ostracods (Crustacea:Ostracoda) in a coastal sand-dune lake,
Loch of Strathbeg, north-east Scotland. Freshwater Biology
22(2):309 – 321.
Bergold, A. 1910. Beiträge zur Kenntnis des innern Baues der
Süßasserostracoden. Zoologische Jahrbücher. Abteilung für
Anatomie und Ontogenie der Tiere 30:1 – 42.
Bernecker, A. 1909. Zur Histologie der Respirationsorgane bei Crustaceen. Zoologischer Jahrbücher. Abteilung für Anatomie und
Ontogenie der Tierecher Abteilung der Anatomie und Ontogenie der Tiere 27:583 – 630.
Bigelow, N. K. 1924. The food of young suckers in Lake Nipigon. University of Toronto Studies, Biological Series Number 24:81 – 116.
Bodergat, A. 1978. L’intensité lumineuse, son influence sur la teneur
en phosphore des carapace d’ostracodes. Géobios 11:715 – 735.
Bowman, T. E., Abele, L. G. 1982. Classification of the recent Crustacea, in: Abele, L. G., Ed., The biology of Crustacea. Vol.
1:systematics, the fossil record, and biogeography. Academic
Press, New York, pp. 1 – 27.
Bronnvall, A. M., Larsson, J. I. R. 1994. Flabelliforma ostracodae n.
sp. (Microspora, Duboscqidae), a new microsporidian parasite
of Candona sp. (Crustacea, Ostracoda) European Journal of
Protistology 30(3):280 – 287.
Bronnvall, A. M., Larsson, J. I. R. 1995. Description of Binucleospora elongata gen. et sp. nov. (Microspora, Caudosporidae), a
microsporidian parasite of ostracods of the genus Candona
(Crustacea, Cyprididae) in Sweden. European Journal of Protistology 31(1):63 – 72.
Broodbakker, N. W. 1983. The genus Strandesia and other
cypricercini (Crustacea, Ostracoda) in the West Indies. Part 1:
Taxonomy. Bijdragen tot de Dierkunde 53:327 – 368.
Broodbakker, N. W., Danielopol, D. L. 1982. The chaetotaxy of
Cypridacea (Crustacea, Ostracoda) limbs; proposals for a descriptive model. Bijdragen tot de Dierkunde 52:103 – 120.
Campbell, C. E. 1995. The influence of a predatory ostracod, Australocypris insularis, on zooplankton abundance and species
composition in a saline lake. Hydrobiologia 302(3):229 – 239.
Cannon, H. G. 1925. On the segmental excretory organs of certain
freshwater ostracods. Philosophical Transactions of the Royal
Society of London 214:1 – 27.
Chaplin, J. A. 1993. The local displacement of a sexually reproducing
ostracod by a conspecific parthenogen. Heredity 71(3):259 – 268.
Chaplin, J. A., Ayre, D. J. 1989. Genetic evidence of variation in the
contributions of sexual and asexual reproduction to populations of the freshwater ostracod Candonocypris novaezelandiae.
Freshwater Biology 22(2):275 – 284.
Chaplin, J. A., Ayre, D. J. 1997. Genetic evidence of widespread dispersal in a parthenogenetic freshwater ostracod. Heredity
78(1):57 – 67.
20. Ostracoda
Chaplin, J. A., Hebert, P. 1997. Cyprinotus incongruens (Ostracoda):an ancient asexual? Molecular Ecology 6(2):155 – 168.
Chaplin, J. A., Havel, J. E., Hebert, P. D. N. 1994. Sex and ostracods
(Review). Trends in Ecology and Evolution 9(11):435 – 439.
Chivas, A. R., DeDeckker, P., Shelley, J. M. G. 1983. Magnesium,
strontium and barium partitioning in nonmarine ostracode
shells and their use in paleoenvironmental reconstruction — a
preliminary study, in: Maddocks, R. F., Ed., Applications of Ostracoda. University of Houston, Department of Geoscience,
Houston, TX, pp. 238 – 249.
Chivas, A. R., DeDeckker, P., Shelley, J. M. G. 1985. Strontium content of ostracods indicate lacustrine paleosalinity. Nature (London) 316:251 – 253.
Chivas, A. R., DeDeckker, P., Shelley, J. M. G. 1986a. Magnesium and
strontium in non-marine ostracod shells as indicators of paleosalinity and palaeotemperature. Hydrobiologia 143:135 – 142.
Chivas, A. R., DeDeckker, P., Shelley, J. M. G. 1986b. Magnesium
content of non-marine ostracod shells:a new paleosalinometer
and palaeothermometer. Palaeogeography, Palaeoclimatology,
Palaeoecology 54:43 – 61.
Creuzé des Châtelliers, M., Marmonier, P. 1990. Macrodistribution
of Ostracoda and Cladocera in a by-passed channel:exchange
between superficial and interstitial layers. Stygologia 5(1):
17 – 24.
Creuzé des Châtelliers, M., Marmonier, P. 1993. Ecology of benthic
and interstitial ostracods (Crustacea) of the Rhône River,
France. Journal of Crustacean Biology 13(2):268 – 279.
Curry, B. B. 1999. An environmental tolerance index for ostracodes
as indicators of physical and chemical factors in aquatic
habitats. Paleogeography, Palaeoclimatology, Palaeoecology
148:51 – 63.
Cvancara, A. M., Clayton, L., Bickley, W. B., Jr., Jacob, A. F.,
Ashworth, A. C., Brophy, J. A., Shay, C. I., Delorme, L. D.,
Lammers, G. E. 1971. Paleolimnology of late Quaternary
deposits, Seibold site, North Dakota. Science 171:172 – 174.
Danforth, W. A. 1948. A list of Iowa ostracods with descriptions of
three new species. Proceedings of the Iowa Academy of Science
55:351 – 359.
Danielopol, D. L. 1969. Recherches sur la morphologie de l’organe
copulateur male chez quelques ostracodes du genre Candona
Baird (Fam. Cyprididae Baird), in: Neale, J. W., Ed., The taxonomy, morphology and ecology of Recent Ostracoda. Oliver &
Boyd, Edinburgh, pp. 136 – 153.
Danielopol, D. L. 1980a. An essay to assess the age of the freshwater
interstitial ostracods of Europe. Bijdragen tot de Dierkunde
50:243 – 291.
Danielopol, D. L. 1980b. Sur la biologie de quelques ostracodes Candoninae épigés et hypogés d’Europe. Bulletin du Musée National d’Histoire Naturelle (Paris) 2:471 – 506.
Danielopol, D. L., Marmonier, P., Boulton, A. J., Bonaduce, G. 1994.
World subterranean ostracod biogeography; dispersal or vicariance. Hydrobiologia 287:119 – 129.
De Deckker, P. 1981. Ostracods of athalassic saline lakes. Hydrobiologia 81:131 – 144.
De Deckker, P. 1983a. Notes on the ecology and distribution of nonmarine ostracods in Australia. Hydrobiologia 106(3):223 – 234.
De Deckker, P. 1983b. Australian Salt Lakes; their history, chemistry,
and biota. Hydrobiologia 105:231 – 244.
De Deckker, P. 1988. The use of ostracods in palaeolimnology in
Australia. Palaeogeography, Palaeoclimatology, Palaeoecology
62:463 – 475.
De Deckker, P., Forester, R. M. 1988. The use of ostracods to reconstruct continental paleoenvironmental records, in DeDeckker,
P., Collin, J.-P., Peypouquet, J.-P., Eds., Ostracoda in the earth
sciences. Elsevier, Amsterdam, pp. 175 – 199.
843
Delorme, L. D. 1967. Field key and methods of collecting freshwater
ostracodes in Canada. Canadian Journal of Zoology 45:
1275 – 1281.
Delorme, L. D. 1968. Pleistocene freshwater Ostracoda from Yukon,
Canada. Canadian Journal of Zoology 46:859 – 876.
Delorme, L. D. 1969a. On the identity of the ostracode genera Cypriconcha and Megalocypris. Canadian Journal of Zoology
47:271 – 281.
Delorme, L. D. 1969b. Ostracodes as Quaternary paleoecological indicators. Canadian Journal of Earth Sciences 6:1471 – 1476.
Delorme, L. D. 1970a. Freshwater ostracodes of Canada. Part I:Subfamily Cypridinae. Canadian Journal of Zoology 48:153 – 169.
Delorme, L. D. 1970b. Freshwater ostracodes of Canada. Part II:Subfamilies Cypridopsinae, Herpetocypridinae, and family Cyclocyprididae. Canadian Journal of Zoology 48:253 – 266.
Delorme, L. D. 1970c. Freshwater ostracodes of Canada. Part III:Family Candonidae. Canadian Journal of Zoology 48:1099 – 1127.
Delorme, L. D. 1970d. Freshwater ostracodes of Canada. Part
IV:Families Ilyocyprididae, Notodromadidae, Darwinulidae,
Cytherideidae, and Entocytheridae. Canadian Journal of Zoology 48:1251 – 1259.
Delorme, L. D. 1971a. Freshwater ostracodes of Canada. Part
V:Families Limnocytheridae, Loxoconchidae. Canadian Journal
of Zoology 49:43 – 64.
Delorme, L. D. 1971b. Paleoecological determinations using Pleistocene freshwater ostracodes. Bulletin du Centre de Recherche
de Pau-SNPA 5:341 – 347.
Delorme, L. D. 1971c. Paleoecology of Holocene sediments from
Manitoba using freshwater ostracodes. Geological Association
of Canada Symposium, Special Paper No. 9:301 – 304.
Delorme, L. D. 1972. Groundwater flow systems, past and present,
in 24th International Geological Congress, Montreal. Section
11, pp. 222 – 226.
Delorme, L. D. 1978. Distribution of freshwater ostracodes in Lake
Erie. Journal of Great Lakes Research 4:216 – 220.
Delorme, L. D. 1982. Lake Erie oxygen, the prehistoric record. Journal of Fisheries and Aquatic Sciences 39:1021 – 1029.
Delorme, L. D. 1989. Methods in Quaternary ecology No. 7:Freshwater Ostracoda. Geosciences Canada 16:85 – 90.
Delorme, L. D. 1996. Burlington Bay, Lake Ontario, its paleolimnology based on fossil ostracodes. Water Quality Research Journal
of Canada 31:643 – 671.
Delorme, L. D., Donald, D. 1969. Torpidity of freshwater ostracodes. Canadian Journal of Zoology 47:997 – 999.
Delorme, L. D., Zoltai, S. C. 1984. Distribution of an arctic ostracod
fauna in space and time. Quaternary Research 21:65 – 73.
Delorme, L. D., Zoltai, S. C., Kalas, L. L. 1977. Freshwater shelled
invertebrate indicators of paleoclimate in Northwestern Canada
during the late glacial times. Canadian Journal of Earth Sciences
14:2029 – 2046.
Delorme, L. D., Zoltai, S. C., Kalas, L. L. 1978. Freshwater shelled
invertebrate indicators of paleoclimate in Northwestern Canada
during late glacial times. Reply. Canadian Journal of Earth Sciences 15:462 – 463.
Deschiens, R. 1954. Mécansime de l’action léthale de Cypridopsis
hartwigi sur les mollusques vecteurs des bilharzioses. Bulletin de
la Société de Pathologie exotique 47:399 – 401.
Deschiens, R., Lamy, L., Lamy, H. 1953. Sur un ostracode prédateur
de bulins et de planorbes. Bulletin de la Société de Pathologie
exotique 46:956 – 958.
Dezfuli, B. S. 1996. Cypria reptans (Crustacea:Ostracoda) as an intermediate host of Neoechinorhynchus rutili (Acanthocephla: Eoacanthocephala) in Italy. Journal of Parasitology 82(3):503 – 505.
Diarra, K., Toguebaye, B. S. 1996. Ultrastructure of Nosema stenocypris Diarra and Toguebaye, 1994, a microsporidian parasite
844
L. D. Delorme
of Stenocypris major (Crustacea, Ostracoda, Cyprididae).
Archiv für Protistenkunde 146(3 – 4):363 – 367.
Dieter, C. D., Duffy, W. G., Flake, L. D. 1996. The effect of phorate
on wetland macroinvertebrates. Environmental Toxicology &
Chemistry 15(3):308 – 312.
Dobbin, C. N. 1941. Fresh-water Ostracoda from Washington and
other western localities. University of Washington Publications
in Biology 4:175 – 246.
Douglas, D. J., Healy, B. 1991. The freshwater ostracods of two
quagmires in Co. Louth, Ireland. Internationale Vereinigung für
Theoretische und Angewandte Limnologie, Verhundlungen
24(3):1522 – 1525.
Engstrom, D. R., Nelson, S. R. 1991. Paleosalinity from trace metals
in fossil ostracodes compared with observational records at
Devils Lake, North Dakota, USA. Palaeogeography, Palaeoclimatology, Palaeoecology 83:295 – 312.
Fassbinder, K. 1912. Beiträge zur Kenntnis der Süsswasserostracoden. Zoologische Jahrbücher. Abteilung für Anatomie und Ontogenie der Tiere 32:533 – 576.
Ferguson, E., Jr. 1944. Studies of the seasonal life history of three
species of fresh-water ostracodes. American Midland Naturalist
32:713 – 727.
Ferguson, E., Jr. 1958a. Seasonal life history studies of two species of
freshwater ostracods. Anatomical Records 131:549 – 550.
Ferguson, E., Jr. 1958b. Freshwater ostracods from South Carolina.
The American Midland Naturalist 59:111 – 119.
Ferguson, E., Jr. 1964. Stenocyprinae, a new subfamily of freshwater
cyprid ostracods (Crustacea) with description of a new species
from California. Proceedings of the Biological Society of Washington 77:17 – 24.
Ferguson, E., Jr. 1967. New ostracods from the playa lakes of eastern
New Mexico and western Texas. Transactions of the American
Microscopical Society 86:244 – 250.
Forester, R. M. 1983. The relationship of two lacustrine ostracode
species to solute composition and salinity, implications for paleohydrochemistry. Geology 11:435 – 439.
Forester, R. M. 1986. Determination of the dissolved anion composition of ancient lakes from fossil ostracodes. Geology 14:796 – 799.
Forester, R. M. 1987. Late Quaternary paleoclimate records from lacustrine ostracodes, in: Ruddiman, W. E., Wright, H. E., Eds.,
North American and adjacent oceans during the last deglaciation. DNAG. Geological Society of America Boulder, CO, pp.
261 – 276.
Forester, R. M. 1991. Ostracode assemblages from springs in the
western United States:implications for paleohydrology. Memoir
of the Entomological Society of Canada 155:181 – 201.
Forester, R. M., Brouwers, E. M. 1985. Hydrochemical parameters
governing the occurrence of estuarine and marginal estuarine
ostracodes, an example from central Alaska. Journal of Paleontology 59:344 – 369.
Forester, R. M., Delorme, L. D., Bradbury, J. P. 1987. Mid-Holocene
climate in Northern Minnesota. Quaternary Research 28:
263 – 273.
Forester, R. M., Colman, S. M., Reynolds, R. L., Keigwin, L. D.
1994. Lake Michigan’s late Quaternary limnological and climate history from ostracode, oxygen isotope, and magnetic susceptibility. Journal of Great Lakes Research 20(1):93 – 107.
Forester, R. M., Delorme, L. D., Bradbury, J. P. 1987. Mid-Holocene
climate in Northern Minnesota. Quaternary Research (N.Y.)
28:263 – 273.
Fox, H. M., Taylor, A. E. 1954. Injurious effect of air-saturated water on certain invertebrates. Nature (London) 174:312.
Fox, H. M., Taylor, A. E. 1955. The tolerance of oxygen by aquatic
invertebrates. Proceedings of the Royal Society of London, Series B 143:214 – 225.
Fryer, G. 1957. The food of some freshwater cyclopoid copepods
and its ecological significance. Journal of Animal Ecology 26:
263 – 286.
Furtos, N. C. 1933. The Ostracoda of Ohio. Ohio Biological Survey,
Vol. 5, Bulletin 29:413 – 524.
Furtos, N. C. 1935. Fresh-water Ostracoda from Massachusetts.
Journal of the Washington Academy of Science 25:530 – 544.
Furtos, N. C. 1936a. On the Ostracoda from the cenotes of Yucatan
and vicinity. Carnegie Institution of Washington, Publ. Number
457:89 – 115.
Furtos, N. C. 1936b. Freshwater Ostracoda from Florida and North
Carolina. American Midland Naturalist 17:491 – 522.
Grant, I. F., Egan, E. A., Alexander, M. 1983. Measurement of rate
of grazing of the ostracod Cyprinotus carolinensis on blue-green
algae. Hydrobiologia 106:199 – 208.
Green, J. 1954. A note on the food of Chaetogaster diaphanus. Annals of the Magazine of Natural History 12:842 – 844.
Green, J. 1962. Bile pigment in Eucypris virens (Jurine). Nature
(London) 196:1318 – 1319.
Griffiths, H. I., Evans, J. G. 1994. Infestation of the freshwater ostracod
Cypria ophthalmica [sic] (Jurine) by the Peritrich Nüchterleinella
corneliae Matthes. Archiv für Protistenkunde 144(3):315–317.
Griffiths, H. I., Martin, D. S., Shine, A. J., Evans, J. G. 1993. The ostracod fauna (Crustacea, Ostracoda) of the profundal benthos
of Loch Ness. Hydrobiologia 254(2):111 – 117.
Griffiths, H. I., Pietrzeniuk, E., Fuhrmann, R., Lennon, J. J.,
Martens, K., Evans, J. G. 1998. Tonnacypris glacialis (Ostracoda, Cyprididae):taxonomic position, (palaeo)ecology, and
zoogeography. Journal of Biogeography 25(3):515 – 526.
Gutentag, E. D., Benson, R. H. 1962. Neogene (Plio-Pleistocene)
fresh-water ostracodes from the central high plains. Bulletin of
the Geological Survey of Kansas. Article 157:60 p.
Hanström, B. 1924. Beiträge zur Kenntnis des zentralen Nervensystem der Ostracoden und Copepoden. Zoologischer Anzeiger
61:31 – 38.
Harding, J. P. 1962. Mungava munda and four other new species of
ostracod crustaceans from fish stomachs. Natural History of the
Rennell Island, British Solomon Islands 4:51 – 62.
Harding, J. P. 1964. Crustacean cuticle with reference to the ostracod
carapace. Pubblicazione della Stazione Zoologica di Napoli
33:9 – 31.
Hart, C. W., Jr., Hobbs, H. H., Jr. 1961. Eight new troglobitic ostracods of the genus Entocythere (Crustacea, Ostracoda) from the
eastern United States. Proceedings of the Academy of Natural
Sciences of Philadelphia 113:173 – 185.
Hart, D. G., Hart, C. W., Jr. 1974. The ostracod family Entocytheridae. Monograph 18. Academy of Natural Sciences of Philadelphia, 239 p.
Have, A. 1993. Effects of area and patchiness on species richness:an
experimental archipelago of ciliate microcosms. Oikos 66(3):
493 – 500.
Havel, J. E., Hebert, P. D. N. 1993. Clonal diversity in parthenogenetic ostracods, in: McKenzie, K. G., Jones, P. J., Eds., Ostracoda in the Earth and Life Sciences. Proceedings of the 11th International, Symposium on Ostracoda. Balkema, Rotterdam,
pp. 353 – 368.
Havel, J. E., Talbott, B. L. 1995. Life history characteristics of the
freshwater ostracod Cyprinotus incongruens and their application to toxicity. Ecotoxicology 4:206 – 218.
Havel, J. E., Hebert, P. D. N., Delorme, L. D. 1990a. Genetics of sexual Ostracoda from a low arctic site. Journal of Evolutionary
Biology 3:65 – 84.
Havel, J. E., Hebert, P. D. N., Delorme, L. D. 1990b. Genotypic diversity of asexual Ostracoda from a low arctic site. Journal of
Evolutionary Biology 3:391 – 410.
20. Ostracoda
Hoff, C. C. 1942. The ostracods of Illinois, their biology and taxonomy. Illinois Biological Monograph 19:196 p.
Hoff, C. C. 1943a. Seasonal changes in ostracod fauna of temporary
ponds. Ecology 24:116 – 118.
Hoff, C. C. 1943b. Two new ostracods of the genus Entocythere and
records of previously described species. Journal of the Washington Academy of Science 33:276 – 286.
Horne, F. R. 1993. Survival strategy to escape desiccation in a freshwater ostracod. Crustaceana 65(1):53 – 61.
Hu, F. S., Ito, E., Brubaker, L. B., Anderson, P. M. 1998. Ostracode
geochemical record of Holocene climatic change and implications for vegetational response in the northwestern Alaska
Range. Quaternary Research 49:86 – 95.
Hummon, W. D. 1981. Extraction by sieving: a biased procedure in
studies of stream meibenthos. Transactions of the American
Microscopical Society 100:278 – 284.
Hurtubise, R. D., Havel, J. E., Little, E. E. 1998. The effects of ultraviolet-B radiation on freshwater invertebrates:experiments with a
solar simulator. Limnology & Oceanography 43(6):1082 – 1088.
Jahn, A., Gamenick, I., Theede, H. 1996. Physiological adaptations
of Cyprideis torosa (Crustacea, Ostracoda) to hydrogen
sulphide. Marine Ecology Progress Series 142(1 – 3):215 – 223.
Johansen, F. 1912. Freshwater life in north-east Greenland. Meddelelser om Gronland 45:321 – 337.
Karrow, P. F., Anderson, T. W., Delorme, L. D., Clarke, A. J., Jr.
1975. Stratigraphy, paleontology, and age of Lake Algonquin
sediments in southwestern Ontario, Canada. Quaternary Research (N.Y.) 5:49 – 87.
Karrow, P. F., Anderson, T. W., Delorme, L. D., Miller, B. B.,
Chapman, L. J. 1995. Late-glacial paleoenvironment of Lake
Algonquin sediments near Clarksburg Ontario. Journal of
Paleolimnology 14:297 – 309.
Kesling, R. V. 1951a. The morphology of ostracod molt stages.
Illinois Biological Monographs 21: 1 – 324.
Kesling, R. V. 1951b. Terminology of ostracod carapaces. Contribution from the Museum of Paleontology, University of Michigan
9:93 – 171.
Kesling, R. V. 1953. A slide rule for the determination of instars in
ostracod species. Contributions from the Museum of Paleontology, University of Michigan 11:97 – 109.
Kesling, R. V. 1957. Notes on Zenker’s organs in the ostracod Candona. American Midland Naturalist 57:175 – 182.
Kesling, R. V. 1965. Anatomy and dimorphism of adult Candona
suburbana Hoff. Four Reports of ostracod Investigations. Report 1. Univ. of Michigan Press, Ann Arbor. 56 p.
King, J. L., Simovich, M. A., Brusca, R. C. 1996. Species richness,
endemism and ecology of crustacean assemblages in northern
California vernal ponds. Hydrobiologia 328:85 – 116.
Klassen, R. W., Delorme, L. D., Mott, R. J. 1967. Geology and paleontology of Pleistocene deposits in southwestern Manitoba.
Canadian Journal of Earth Sciences 4:433 – 447.
Klie, W. 1931. Campagne spéologique de C. Bolivar et R. Jeannel
dan L’Amérique du Nord (1928). Part 3: Crustacés Ostracodes.
Archives Zoologie Expérimentale Générale 71:333 – 344.
Klie, W. 1939. Süsswasserostracoden aus Nordostbrasilien. Part 1.
Zoologischer Anzeiger 128:84 – 91.
Klugh, A. B. 1927. The ecology, food-relations and culture of freshwater Entomostraca. Transactions of the Royal Canadian Institute 16:15 – 98.
Kornicker, L. S., Sohn, I. G. 1971. Viability of ostracode eggs egested
by fish and effect of digestive fluids on ostracode shells; ecologic
and paleoecologic implications, in: Oertli, H. J., Ed., Paleoecologie des ostracodes. Bulletin du Centre de Recherches de
Pau-SNPA, Supplement 5:125 – 135.
Korschelt, E. 1915. Über das Verhalten verschiedener wirbelloser
845
Tiere gegen niedere Temperaturen. Zoologischer Anzeiger
45:113 – 115.
Külköylüoĝlu, O., Vinyard, G. L. 1998. A new bisexual form of Cavernocypris subterranea (Wolf, 1920) (Crustacea, Ostracoda)
from Idaho. Great Basin Naturalist. 58(4):380 – 385.
Lamb, H. F., Gasse, F., Benkaddour, A., El Hamouti, N., van der
Kaars, S., Perkins, W. T., Pearce, N. J., Roberts, C. N. 1995.
Relation between century-scale Holocene arid intervals in tropical and temperate zones. Nature 373(6510):134 – 137.
Lancaster, J., Robertson, A. L. 1995. Microcrustacean prey and
macroinvertebrate predators in a stream food web. Freshwater
Biology 34(1):123 – 134.
Landis, W. G., Matthews, G. B., Matthews, R. A., Sergeant, A. 1994.
Application of multivariate techniques to endpoint determination, selection and evaluation in ecological assessment. Environmental Toxicology and Chemistry 13(2):1917 – 1927.
Larsen, J., Birks, H. J. H., Raddum, G. G., Fjellheim, A. 1996.
Quantitative relationships of invertebrates to pH in Norwegian
river systems. Hydrobiologia 328(1):57 – 74.
Lécher, P., Defaye, D., Noel, P. 1995. Chromosomes and nuclear
DNA of Crustacea. Invertebrate Reproduction and Development 27(2):85 – 114.
Little, T. J., Hebert, P. D. N. 1994. Abundant asexuality in tropical
freshwater ostracodes. Heredity 73:549 – 555.
Little, T. J., Hebert, P. D. N. 1996. Endemism and ecological islands:
the ostracods from Jamaican bromeliads. Freshwater Biology
36(2):327 – 338.
Lowndes, A. G. 1935. The sperms of fresh-water ostracods. Proceedings of the Zoological Society of London. Part 2:35 – 48.
Maddocks, R. F. 1982. Ostracoda, in: Abele, L. G., Ed., The biology
of Crustacea, Vol. 1:Systematics, the fossil record, and biogeography. Academic Press, New York, pp. 221 – 239.
Maher, L. J., Miller, N. G., Baker, R. G., Curry, B. B., Mickelson, D.
M. 1998. Paleobiology of the sand beneath the Valders diamicton at Valders, Wisconsin. Quaternary Research 49:208 – 221.
Malmquist, B., Meisch, C., Nilsson, A. N. 1997. Distribution patterns of freshwater Ostracoda (Crustacea) in the Canary Islands
with regards to habitat use and biogeography. Hydrobiologia
347:159 – 170.
Marmonier, P., Creuzé des Châtelliers, M. 1991. Effects of spates on
interstitial assemblages of the Rhône River, importance of spatial heterogeneity. Hydrobiologia 210(3):243 – 251.
Marmonier, P., Creuzé des Châtelliers, M. 1992. Biogeography of the
benthic and interstitial living ostracods (Crustacea) of the
Rhône River (France). Journal of Biogeography 19:693 – 704.
Marmonier, P., Ward, J. V. 1990. Superficial and interstitial Ostracoda of the South Platt River (Colorado, USA) — systematics
and biogeography. Stygologia 5(4):225 – 239.
Marmonier, P., Bodergat, A. M., Doledec, S. 1994. Theoretical habitat templets, species traits, and species richness:ostracods (Crustacea) in the upper Rhône River and its floodplain. Freshwater
Biology 31(3):341 – 355.
Marmonier, P., Meisch, C., Danielopol, D. L. 1989. A review of the
Genus Cavernocypris Hartmann (Ostracoda, Cypridopsinae):
Systematics, Ecology and Biogeography. Bulletin de la Société de
Luxembourg 89:221 – 278.
Marshall, W. S. 1903. Entocythere cambaria (nov. gen. et nov. spec.),
a parasitic ostracod. Transactions of the Wisconsin Academy of
Science, Arts, and Letters 14:117 – 144.
Martens, K. 1986. Taxonomic revision of the subfamily Megalocypridinae Rome, 1965. Verhandelingen van de Koninklijke
Academie voor Wetenscappen, Letteren en schone Kunsten van
Belgie 48:1 – 81.
Martens, K. 1989. On the systematic position of the Eucypris
clavata — group, with a description of Trajancypris gen. nov.
846
L. D. Delorme
(Crustacea, Ostracoda). Archiv für Hydrobiologie, Supplement
83(2):227 – 251.
Martens, K., de Moor, F. 1995. The fate of the rhino ridge pool
at Thomas Baines Nature Reserve:cautionary tale for nature
conservationists. South African Journal of Science 91(8):
385 – 387.
Martens, K., DeDeckker, P., Marples, T. G. 1985. Life history of
Mytilocypris henricae (Chapman)(Crustacea:Ostracoda) in Lake
Bathurst, New South Wales. Australian Journal of Marine and
Freshwater Research 36:807 – 819.
Mbahinzireki, G., Uiblein, F., Winkler, H. 1991. Microhabitat selection of ostracods in relation to predation and food. Hydrobiologia 222(2):115 – 119.
McAllister, D. E., Harrington, C. R. 1969. Pleistocene grayling, Thymallus, from Yukon, Canada. Canadian Journal of Earth Sciences 6:1185 – 1190.
McGregor, D. L. 1967. Rhythmic pulsation of the hepatopancreas in
freshwater ostracods. Transactions of the American Microscopical Society 86:166 – 169.
McGregor, D. L. 1969. The reproductive potential, life history and
parasitism of the freshwater ostracod, Darwinula stevensoni
(Brady & Roberstson), in: Neale, J. W., Ed., Taxonomy, morphology and ecology of recent Ostracoda. Oliver & Boyd, Edinburgh, pp. 194 – 221.
McGregor, D. L., Kesling, R. V. 1969a. Copulatory adaptations in
ostracods. Part 1:Hemipenes of Candona. Contributions to the
Museum of Paleontology University of Michigan 22:169 – 191.
McGregor, D. L., Kesling, R. V. 1969b. Copulatory adaptations in
ostracods. Part 2:Adaptations in living ostracods. Contributions
to the Museum of Paleontology University of Michigan
22:221 – 239.
McKenzie, K. G. 1977. Illustrated generic key to South African continental Ostracoda. Annals of the South African Museum
74(3):45 – 103.
McKillop, W. B., Patterson, R. T., Delorme, L. D., Norgrady, T.
1992. The origin, physico-chemistry and biotics of sodium
chloride dominated saline waters on the western shore of
Lake Winnipegosis, Manitoba. Canadian Field Naturalist 106:
454 – 473.
McLay, C. L. 1978a. Comparative observations on the ecology of
four species of ostracods living in a temporary freshwater puddle. Canadian Journal of Zoology 56:663 – 675.
McLay, C. L. 1978b. The population biology of Cyprinotus carolinensis and Herpetocypris reptans (Crustacea, Ostracoda).
Canadian Journal of Zoology 56:1170 – 1179.
McLay, C. L. 1978c. Competition, coexistence, and survival; a computer simulation study of ostracodes living in a temporary puddle. Canadian Journal of Zoology 56:1744 – 1758.
Meisch, C. 1996. Contribution to the taxonomy of Pseudocandona
and four related genera, with the description of Schellencandona
nov. gen., a list of the Candoninae genera, and a key to the European genera of the subfamily (Crustacea, Ostracoda). Bulletin de
la Société des Naturalistes luxembourgeois 97:211 – 237.
Moore, R. C., Ed. 1961. Treatise on invertebrate paleontology. Part
Q:Arthropoda 3, Crustacea, Ostracoda. Geological Society of
America, Univ. of Kansas Press. Lawrence 442 pp.
Müller, G. W. 1912. Ostracoda, in: Schulze, F. E., Ed. Das Tierreich.
Friedländer und Sohn, Lief, Berlin. 434 pp.
Müller-Cale, K. 1913. Über die Entwicklung von Cypris incongruens.
Zoologische Jahrbücher. Abteilund für Anatomie und Ontogenie der Tiere 36:1 – 56.
Nalepa, T. F., Robertson, A. 1981. Screen mesh size affects estimates
of macro- and meio-benthos abundance and biomass in the
Great Lakes. Canadian Journal of Fisheries and Aquatic Sciences 38:1027 – 1034.
Neale, J. W. 1988. Ostracods and paleosalinity reconstruction, in:
DeDeckker, P., Colin, J.-P., Peypouquet, J.-P., Eds., Ostracoda in
the Earth Sciences. Elsevier, Amsterdam, pp. 125 – 155.
Neale, J. W., Delorme, L. D. 1985. Cytheromorpha fuscata, relict
Holocene marine ostracode from freshwater inland lakes of
Manitoba, Canada. Revista Espanola de Micropaleontologia
17:41 – 64.
Newrkla, P. 1985. Respiration of Cytherissa lacustris (Ostracoda) at
different temperatures and its tolerance towards temperature
and oxygen concentration. Oecologia 67:250 – 254.
Novikoff, M. 1908. Über den Bau des Midiamauges der Ostracoden.
Zeitschrift für wissenschaftliche Zoologie 91:81 – 92.
Nudds, T. D., Bowlby, J. N. 1984. Predator-prey size relationships in
North American dabbling ducks. Canadian Journal of Zoology
62(10):2002 – 2008.
Oertli, B. 1995. Spatial and temporal distribution of the zoobenthos
community in a woodland pond (Switzerland). Hydrobiologia
300 / 301:195 – 204.
Peck, S. G. 1994. Diversity and zoogeography of the non-oceanic
Crustacea of the Galápagos Islands, Ecuador (excluding terrestrial Isopoda). Canadian Journal of Zoology 72:54 – 69.
Porter, S. C., Sauchyn, D. J., Delorme, L. D. 1999. The ostracode
record from Harris lake, southwestern Saskatchewan, 9200
years of local environmental change. Journal of Paleolimnology
21:35 – 44.
Proctor, V. W. 1964. Viability of crustacean eggs recovered from
ducks. Ecology 45:656 – 658.
Przibram, H. 1931. Connecting laws in animal morphology. Four
lectures held at the University of London, March, 1929. Univ.
of London Press, London. 62 pp.
Purper, I., Würdig-Macial, N. I. 1974. Occurrence of Heterocypris
incongruens (Ramdohr), 1808 — Ostracoda — in Rio Grande do
Sul, Brazil; discussion on the allied genera Cyprinotus, Hemicypris, Homocypris and Eucypris. Pesquisas 3:69 – 91.
Ranta, E. 1979, Population biology of Darwinula stevensoni (Crustacea, Ostracoda) in an oligotrophic lake. Annales Zoologici
Fennici 16:28 – 35.
Reyment, R. A., Brännström, B. 1962. Certain aspects of the physiology of Cypridopsis (Ostracoda, Crustacea). Acta Universitatis
Stockholmiensis 9:207 – 242.
Rieradevall, M., Roca, J. R. 1995. Distribution and population dynamics of ostracodes (Crustacea, Ostracoda) in a karstic lake: Lake
Banyoles (Catalonia, Spain). Hydrobiologia 310(3): 189 – 196.
Roback, S. S. 1969. Notes on the food of Tanypodine larvae. Entomological News 80:13 – 19.
Roca, J. R., Wansard, G. 1997. Temperature influence on development and calcification of Herpetocypris brevicaudata Kaufmann, 1900 (Crustacea:Ostracoda) under experimental conditions. Hydrobiologia 347:91 – 95.
Roca, J. R., Baltanas, A., Uiblein, F. 1993. Adaptive responses in
Cypridopsis vidua (Crustacea, Ostracoda) to food and shelter
offered by a macrophyte (Chara fragilis). Hydrobiologia
262(2):127 – 131.
Rogulj, B., Marmonier, P., Lattinger, R., Danielopol, D. 1994. Finescale distribution of hypogean Ostracoda in the interstitial habitats of the Rivers Sava and Rhone. Hydrobiologia 287(1):19 – 28.
Rossi, V., Menozzi, P. 1990. The clonal ecology of Heterocypris incongruens (Ostracoda). Oikos 57(3):388 – 398.
Rossi, V., Gandolfi, A., Menozzi, P. 1996. Egg diapause and clonal
structure in parthenogenetic populations of Heterocypris incongruens (Ostracoda). Hydrobiologia 320(1 – 3):45 – 54.
Rossi, V., Rozzi, M. C., Menozzi, P. 1991a. Life strategy differences
among electrophoretic clones of Heterocypris incongruens
(Crustacea, Ostracoda). Verhanlungen internationale Verheinlungen Limnologische 24:2416 – 2819.
20. Ostracoda
Rossi, V., Schön, I., Butlin, R. K., Menozzi, P. 1998. Clonal genetic
diversity, in: Martens, K., Ed., Sex and Parthenogenesis —
evolutionary ecology of reproductive modes in non-marine
ostracods. Backhuys Publ., Leiden, pp. 257 – 274.
Rozzi, M. C., Rossi, V., Benassi, G., Menozzi, P. 1991. Effetto della
temperatura e del fotoperiodo sull’attivazione di uova durature
di dloni elettroforetici di Heterocypris incongruens (Ostracoda).
Società Italieana di Ecologia Atti 12:591 – 594.
Rome, D. R. 1947. Herpetocypris reptans (Ostracode) Étude morphologique et histologique I, morphologie externe et système
nerveux. Cellule 51:51 – 152.
Rome, D. R. 1969. Morphologie de l’attache de la furca chez les
Cyprididae et son utilisation en systematique, in: Neale, J. W.,
Ed., The taxonomy, morphology and ecology of recent Ostracoda. Oliver & Boyd, Edinburgh, pp. 168 – 193.
Rouch, R., Danielopol, D. L. 1997. Species richness of microcrustacea
in subterranean freshwater habitats: comparative analysis and
approximate evaluation. Internationale Revue der Gesamten
Hydrobiologie 82(2):121 – 145.
Sanders, H. O. 1970. Toxicities of some herbicides to six species of
freshwater crustaceans. Journal of Water Pollution Control Federation. 42:1544 – 1550.
Särkkä, J., Levonen, L., Mäkelä, J. 1997. Meiofauna of springs in
Finland in relation to environmental factors. Hydrobiologia
347:139 – 150.
Sars, G. O. 1898. On Megalocypris princeps, a gigantic freshwater
ostracod from South Africa. Archiv for Mathematik og Naturvidenskab 20:1 – 18.
Sars, G. O. 1901. Contributions to the knowledge of the fresh-water
Entomostraca of South America, as shown by artificial hatching
of dried material. Archiv for Mathematik og Naturvidenskab
24:16 – 52.
Sars, G. O. 1926. Freshwater Ostracoda from Canada and Alaska.
Report of the Canadian Arctic Expedition 1913 – 1918 7:3 – 23.
Sars, G. O. 1928. An account of the Crustacea of Norway. Bergen
Museum, Bergen. 277 pp.
Schleip, W. 1909. Vergleichende Untersuchung der Eireifung bei
parthenogenetisch und bei geschlechtich sich fortplanzendan
Ostracoden. Archiv für Zellforschung 2:390 – 431.
Schön, I., Butlin, R. K., Griffiths, H. I., Martens, K. 1998. Slow molecular evolution in an ancient asexual ostracod. Proceedings of
the Royal Society, London B 265:235 – 242.
Schram, F. R. 1986. Crustacea. Oxford Univ. Press, New York. 606 pp.
Schreiber, E. 1922. Beiträge zur Kenntnis der Morphologie, Entwicklung und Lebensweise der Süsswasser-Ostracoden. Zoologische
Jahrbücher. Abteilung für Anatomie und Ontogenie der Tiere.
43:485 – 539.
Sharpe, R. W. 1903. Report of the fresh-water Ostracoda of the
United States National Museum including a revision of the
subfamilies and genera of the family Cyprididae. Proceedings of
the U.S. National Museum. 26:969 – 1001.
Sharpe, R. W. 1918. The Ostracoda, in: Ward, H. B., Whipple, G. C.,
Eds., Fresh-water biology. Wiley, New York, pp. 790 – 827.
Smith, A. J., Donovon, J. J., Ito, E., Engstrom, D. R. 1997. Groundwater processes controlling a prairie lake’s response to middle
Holocene drought. Geology 25(5):391 – 394.
Smith, A. J., Delorme, L. D., Forester, R. M. 1992. A lakes’s solute
history from ostracodes: comparison of methods, in: Kharaka,
Y. K., Maest, A. S., Eds., Water-Rock Interaction, Balkema,
Rotterdam, pp. 677 – 680.
Smith, R. N. 1965. Musculature and muscle scars of Chlamydotheca
arcuata (Sars) and Cypridopsis vidua (O. F. Müller) (Ostracoda:
Cyprididae), Vol. 3, Four Reports on ostracode investigations.
University of Michigan Press, Ann Arbor, MI, pp. 1 – 40.
Sohn, I. G., Kornicker, L. S. 1975. Variation in predation behavior
847
of ostracode species on Schistosmoiasis vector snails. Bulletin of
American Paleontologist. 65:217 – 223.
Sohn, I. G., Kornicker, L. S. 1979. Viability of freeze-dried eggs of
the freshwater Heterocypris incongruens, in: Proceedings of the
Seventh International Symposium on Ostracoda, Belgrade,
pp. 1 – 3.
Staplin, F. L. 1963a. Pleistocene Ostracoda of Illinois. Part 1: Subfamilies Candoninae, Cyprinae, general ecology, morphology.
Journal of Paleontology 37:758 – 797.
Staplin, F. L. 1963b. Pleistocene Ostracoda of Illinois. Part 2: Subfamilies Cyclocyprinae, Cypridopsinae, Ilyocyprinae; families
Darwinulidae and Cytheridae, stratigraphic ranges and assemblage patterns. Journal of Paleontology 37:1164 – 1203.
Strayer, D. 1985. The benthic micrometazoans of Mirror Lake,
New Hampshire. Archiv für Hydrobiologie, Supplement 72:
287 – 426.
Strayer, D. 1988. Life history of a lacustrine ostracod. Hydrobiologia
160(2):189 – 191.
Swain, F. M. 1963. Pleistocene Ostracoda from the Gubik formation,
Arctic Coastal Plain, Alaska. Journal of Paleontology 37:
798 – 834.
Swüste, H. F. J., Cremer, R., Parma, S. 1973. Selective predation by
larvae of Chaoborus flavicans (Diptera, Chaoboridae). Internationale Vereinigung für Theoretische und Angewandte Limnologie. Verhundlungen 18:1559 – 1563.
Sylvester-Bradley, P. C. 1941. The shell structures of the Ostracoda
and its application to their palaeontological investigation. The
Annals and Magazine of Natural History Ser. 11:1 – 33.
Sylvester-Bradley, P. C., Benson, R. H. 1971. Terminology for surface
features in ornate ostracodes. Lethaia 4:249 – 286.
Sywula, T., Glazewska, I., Whatley, R. C., Moguilevsky, A. 1995.
Genetic differentiation in the brackish-water ostracod Cyprideis
torosa. Marine Biology 121(4):647 – 653.
Tabacchi, E., Marmonier, P. 1994. Dynamics of the interstitial ostracod assemblage of a pond in the Adour alluvial plain. Archiv
fur Hydrobiologie 131(3):321 – 340.
Takamura, K., Yasuno, M. 1986. Effects of pesticide application on
chironomid larvae and ostracods in rice fields: Applied Entomology and Zoology 21(3):370 – 376.
Towle, E. W. 1900. A study in the heliotropism of Cypridopsis. The
American Journal of Physiology 3:345 – 365.
Tressler, W. L. 1949. Freshwater Ostracoda from Brazil. Proceedings
of the United States National Museum 100(3258):61 – 83.
Tressler, W. L. 1954. Fresh-water Ostracoda from Texas and Mexico.
Journal of the Washington Academy of Science 44:138 – 149.
Tressler, W. L. 1956. Ostracoda from bromeliads in Jamaica and
Florida. Journal of the Washington Academy of Science 16:
333 – 336.
Tressler, W. L. 1957. The Ostracoda of Great Slave Lake. Journal of
the Washington Academy of Science 47:415 – 423.
Triebel, E. 1953. Genotypus und Schalen-Merkmale der OstracodenGattung Stenocypris. Senckenbergiana 34:5 – 14.
Turgeon, J., Hebert, P. D. N. 1994. Evolutionary interactions
between sexual and all-female taxa of Cyprinotus (Ostracoda,
Cyprididae). Evolution 48(6):1855 – 1865.
Turgeon, J., Hebert, P. D. N. 1995. Genetic characterization of
breeding systems, ploidy levels and species boundaries in
Cypricercus (Ostracoda). Heredity 75(6):561 – 570.
Turner, C. H. 1896. Morphology of the nervous system of Cypris.
Journal of Comparative Neurology 6:20 – 44.
Turpen, J. B., Angell, R. W. 1971. Aspects of molting and calcification in the ostracod Heterocypris. The Biological Bulletin
(Woods Hole, Mass.) 140(2):331 – 338.
Uiblein, F., Roca, J. R., Danielopol, D. L. 1994. Experimental observations on the behavior of the ostracode Cypridopsis vidua.
848
L. D. Delorme
Internationale Vereinigung für Theoretische und Angewandte
Limnologie, Verhundlungen 25:2418 – 2420.
van Harten, D. 1975. Size and environmental salinity in the modern
euryhaline ostracod Cyprideis torosa (Jones, 1850), a biometrical study. Palaeogeography, Palaeoclimatology, Palaeoecology
17(1):35 – 48.
von Grafenstein, V., Erlenkeuser, H., Müller, J., Kleinmann-Eisenmann, A. 1992. Oxygen isotope records of benthic ostracods in
Bavarian Lake sediments. Naturwissenschaften 79:145 – 152.
von Grafenstein, V., Erlenkeuser, H., Kleinmann, A., Müller, J., Trimborn, P. 1994. High-frequency climatic oscillations during the
last deglaciation as revealed by oxygen-isotope records of benthic organisms (Ammersee, southern Germany). Journal of Paleolimnology 11:349 – 357.
Walton, M., Hobbs, H. H., Jr. 1959. Two new eyeless ostracods of
the genus Entocythere from Florida. Quarterly Journal of the
Florida Academy of Science 22:114 – 120.
Ward, J. V., Stanford, J. A., Voelz, N. J. 1994. Spatial distribution
patterns of Crustacea in the floodplain aquifer of an alluvial
river. Hydrobiologia 287:11 – 17.
Westgate, J. A., Delorme, L. D. 1988. Lacustrine ostracodes in the
late Pleistocene Sunnybrook diamicton of southern Ontario,
Canada. Reply. Canadian Journal of Earth Sciences 25:
1717 – 1720.
Westgate, J. A., Chen, F.-J., Delorme, L. D. 1987. Lacustrine ostracodes in the late Pleistocene Sunnybrook diamicton of southern
Ontario. Canadian Journal of Earth Sciences 24:2330 – 2335.
Weygoldt, P. 1960. Embryologische Untersuchungen an Ostrakoden;
Die Entwicklung von Cyprideis litoralis (G. S. Brady) (Ostracoda, Podocopa, Cytheridae). Zoologische Jahrbücher.
Abteilund für Anatomie und Ontogenie der Tiere 78:369 – 426.
Weygoldt, P. 1961. Zur Kenntnis der Sekretion im Zentralnervensystem der Ostrakoden Cyprideis litoralis (G. S. Brady) (Podocopa
Cytheridae) und Cypris pubera (O.F.M.) (Podocopa Cypridae),
Neurosekretion und Sekretionzellen im Perineurium. Zoologischer Anzeiger 166:69 – 70.
White, M. J. D. 1973. Animal cytology and evolution. Cambridge
Univ. Press, London.
Wickstrom, C. E., Castenholz, R. W. 1973. Thermophilic ostracod:
aquatic metazoan with the highest known temperature tolerance. Science 181:1063 – 1064.
Wiggins, G. B., Mackay, R. J., Smith, I. M. 1980. Evolutionary and
ecological strategies of animals in annual temporary ponds.
Archiv für Hydrobiologie Supplement 58:97 – 206.
Wohlgemuth, R. 1914. Beobactungen und Untersuchungen über die
Biologie der Süsswasserostracoden; ihr Vorkommen in Sachsen
und Böhmen, ihr Lebensweise und ihr Fortpflanzung. Biologisches Supplement zur Internationale Revue der Gesamten Hydrobiologie und Hydrobiologie 6:1 – 72.
Woltereck, R. 1898. Zur Bildung und Entwickelung des OstracodenEies. Zeitschrift für Wissenschaftliche Zoologie 64:596 – 620.
Xia, J., Engstrom, D. R., Ito, E. 1997a. Geochemistry of ostracode
calcite, part 2. The effects of water chemistry and seasonal
temperature variation on Candona rawsoni. Geochimica et
Cosmochimica Acta 61(2):383 – 391.
Xia, J., Haskell, B. J., Engstrom, D. R., Ito, E. 1997b. Holocene
climate reconstruction from tandem trace-element and stable
isotope composition of ostracodes from Coldwater Lake, North
Dakota, USA. Journal of Paleolimnology 17:85 – 100.
Xia, J., Ito, E., Engstrom, D. R. 1997c. Geochemistry of ostracode
calcite, part 1. An experimental determination of oxygen isotope fractionation. Geochimica et Cosmochimica Acta 61(2):
377 – 382.
Yu, Z., Ito, E. 1999. Possible solar forcing of century-scale drought
frequency in the northern Great Plains. Geology 29(3):263 – 266.
Zelmer, D. A., Esch, G. E. 1998. Interactions between Halipegus occidualiis and its ostracod second intermediate host: evidence for
castration? Journal of Parasitology 84(4):778 – 782.
Zissler, D. 1969a. Die Spermiohistogenese des SüsswasserOstracoden. Part 1: Die ovalen und spindelförmigen Spermatiden. Zeitschrift für Zellforschung und Mikroscopische
Anatomie 96:87 – 105.
Zissler, D. 1969b. Die Spermiohistogenese SüsswasserOstracoden
Notodromas monacha O. F. Müller. Part 2:Die spindelförmigen
und schlauchürmigen Spermatiden. Zeitschrift für Zellforschung
und Mikroscopische Anatomie 96:106 – 133.
Zissler, D. 1970. Zur Spermiohistogenese im Vas Deferens von
Süsswasser-Ostracoden. Cytobiologie 2:83 – 86.
21
CLADOCERA AND OTHER
BRANCHIOPODA
Stanley I. Dodson
David G. Frey†
Department of Zoology
University of Wisconsin
Madison, Wisconsin 53706
I. Introduction
CLADOCERA
II. Anatomy and Physiology
A. General Form and Function
B. Metabolism
III. Distribution
A. Zoogeography
B. Cladocera and the Species Concept
C. Biodiversity
IV. Behavior
A. Mating Behavior
B. Individual Swimming Behavior
C. Diel Vertical Migration
D. Swarming Behavior
E. Escape Behavior
V. What Cladocerans Eat and What
Eats Cladocerans
A. Cladoceran Feeding
B. Predators on Cladocerans
VI. Population Regulation
VII. Functional Role in the Ecosystem
VIII. Paleolimnology and
Molecular Phylogeny
A. Paleolimnology
B. Cladoceran Fossil Record
C. Molecular Phylogeny
IX. Toxicology
X. Current and Future Research
XI. Collecting and Rearing
A. Sampling Techniques
B. Culture Methods
XII. Taxonomic Keys for Cladoceran Genera
A. Available Keys and
Specimen Preparation
B. Taxonomic Key to Families
of Freshwater Cladocerans
C. Taxonomic Key to Genera
of the Family Holopediidae
D. Taxonomic Key to Genera
of the Family Sididae
E. Taxonomic Key to Genera
of the Family Chydoridae
(North of Mexico)
F. Taxonomic Key to Genera
of the Family Daphniidae
and Moinidae
G. Taxonomic Key to Genera
of the Family Bosminidae
H. Taxonomic Key to Genera of
Superfamily Macrothricoidea
I. Key to the Orders Onychopoda
and Haplopoda
OTHER BRANCHIOPODS
XIII. Anatomy and Physiology of
The Other Branchiopods
A. General External and Internal
Anatomic Features
B. Relevant Physiological
Information
XIV. Ecology of the Other Branchiopods
A. Life History
B. Distribution and Biogeography
C. Physiological Adaptations
D. Behavior
E. Foraging Relationships
F. Population Regulation
G. Functional Role in the
Ecosystem
H. Toxicology
XV. Current and Future Research
Problems
XVI. Collecting and Rearing Techniques
XVII. Taxonomic Keys for Non-Cladoceran
Genera of Branchiopods
A. Available Keys and
Specimen Preparation
B. Taxonomic Key to Genera
of Non-Cladoceran Freshwater
Branchiopoda
Literature Cited
†Dr. David G. Frey’s contribution to this chapter is greatly appreciated, and all of us regret his death prior to its publication.
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
849
850
S. I. Dodson and D. G. Frey
TABLE I The Families and Present Number of Extant
Genera of Class Branchiopoda
I. INTRODUCTION1
Branchiopods are small crustaceans that have flattened leaflike legs. They occur in all freshwater habitats
and can be abundant enough to form conspicuous
swarms. Branchiopods are useful for studies of community and population ecology, animal behavior, functional morphology, evolution of life history. They occupy a key position in aquatic communities, both as
important herbivores eating algae and bacteria and as
major prey items of fish, birds, backswimmers, and
other aquatic predators. Their fossil remains open a
window into the past climate and ecology of lakes.
The orders of branchiopods (Table I), including
eight living and two extinct orders are related to each
other only distantly (Fryer, 1987; Kerfoot and Lynch,
1987; Olesen, 1998). The Branchiopoda are a heterogeneous group of crustaceans that share a few characteristics, principally similar thoracic legs called phyllopods, which are flat, edged with setae, not distinctly
segmented, and usually appear to be unbranched.
While this leg architecture resembles that of some of
the earliest crustacean fossils, it is probable that the
similarities among the branchiopods is due at least as
much to convergent evolution as to common ancestry.
Branchiopods also have similar mouthparts. Mandibles
are simple unsegmented rods with corrugated grinding
inner surfaces. The first and second pairs of maxillae
are reduced to small scalelike structures, or are absent.
Branchiopods have on the last body segment a pair of
spines or claws. A controversy exists concerning the
evolutionary significance of these terminal spines (e.g.,
Bowman, 1971; Schminke, 1976).
Because branchiopods are such a diverse group,
this chapter is divided into two major sections. The
first focuses on the “cladocera,” a group of four orders
of small-sized branchiopods (Freyer 1987, Olesen
1998). The second section focuses on the “non-cladoceran orders,” which are less diverse and typically
somewhat larger than the cladocera.
CLADOCERA
Most plankton ecologists now agree that the term
“cladocera” has no taxonomic significance (because
the orders are probably not closely related), although
1
The references in this chapter are helpful starting places for futher
reading and research. To find our more about a topic of interest,
search electronic biological data bases, such as Biological Abstracts
and the Zoalogical Record, which can be accessed at many university libraries. One typically finds a rich mine of publications. If your
library lacks the publication, many are available through interlibrary loan programs
Order
Family
Number of genera
in world
Anomopoda
Daphniidaea
Moinidaeb
Bosminidaec
Ilyocryptidaed
Macrothricidaee
Neothricidaee
Acantholeberidaee
Ophryoxidaee
Chydoridaef
Sididaeg
Holopediidaeg
Podonidae h
Polyphemidaeh
Cercopagidaeh
Leptodoridaeh
Artemiidae
Branchinectidae
Branchipodidae
Chirocephalidae
Linderiellidae
Polyartemiidae
Streptocephalidae
Thamnocephalidae
Cyclestheriidae
Cyzcidae
Leptestheriidae
Limnadiidae
Lynceidae
Triopsidae
8
2
2
1
10
2
1
2
37
7
1
7
1
2
1
1
1
6
7
2
2
1
1
4
4
3
6
3
2
Ctenopoda
Onychopoda
Haplopoda
Anostracai
Spinicaudatai
Laevicaudataj
Notostracak
The “cladocera” include the first four orders listed in the table. The
“conchostraca” include the orders Spinicaudata and Laevicaudata.
a
Number of genera of Daphniidae, see Fryer (1991).
b
For Moinidae, see Goulden (1968) and Fryer (1991).
c
For Bosminidae, see DeMelo and Hebert (1994).
d
For Ilyocryptidae, see Smirnov (1992).
e
The families Macrothricidae, Neothricidae, Acantholeberidae, and
Ophryoxidae are discussed in Smirmov (1992) and grouped into
the superfamily Macrothricoidea by Dumont and Silva-Briano
(1998).
f
For family Chydoridae; see Smirnov (1974, 1996). Alonso (1996)
suggests a name change to Eurycercidae. Because of long usage, I
prefer the name “Chydoridae” for the family.
g
For Sididae and Holopedidae, see Korovchinsky (1992).
h
For the predaceous cladocerans, see Rivier (1998).
i
Families and number of genera are based on an annotated update
(personal communication) by D. Belk, based on Belk (1982).
j
For Laevicaudata see Martin and Belk (1988).
k
For Notostraca, see Fryer (1988).
the term is still used for convenience. The cladocera are
grouped into four orders (Fryer, 1987), 15 families (including the revisions suggested by Dumont and SilvaBriano, 1998), about 80 genera, and roughly 400
species (Table I). There is some controversy over
whether the orders Onychopoda and Haplopoda
21. Cladocera and Other Branchiopoda
(which are related predaceous species; Olesen, 1998)
are even members of the Branchiopoda (Starobogatov,
1986). The number of genera in each order (Table I) is
only approximate, because new genera have been established recently in the families Sididae, Daphniidae,
Macrothricidae, and Chydoridae, with certainly more
to come.
Almost all members of the suborder Radopoda
(Dumont and Silva-Briano, 1998) and some members
of the Sididae (Table I) are benthic (bottom-dwelling),
living on and in various surfaces, particularly aquatic
plants (macrophytes), coarse plant detritus (decaying
plant material), and organic sediments. They are called
“meiobenthos” (Frey, 1988a) because of their small
size (mostly 1 mm in total length) and because of
their benthic habitat preference. Meiobenthos are often
most abundant in the plants and associated sediments
of the littoral (or shallow water) zone. Chydorid populations commonly have more than a million animals
per square meter of bottom, and twenty or more
species can coexist in the same small bit of habitat.
Most species in the remaining eight families in Table I
are primarily or totally planktonic (inhabiting the open
water zone), where their facility for swimming makes
them quite independent of surfaces.
Besides differing in habitat, planktonic and
meiobenthic cladocerans differ also in ecological relationships and in their evolution. Hence, to imply, as
students of plankton commonly do, that everything
learned about Daphnia or Bosmina applies equally
well to all cladocera is often misleading or incorrect.
The present chapter seeks to present an even treatment
of all cladoceran families.
II. ANATOMY AND PHYSIOLOGY
A. General Form and Function
1. General Female Form and Function
Martin (1992) provides a masterful and detailed
description of branchiopod anatomy, with a wealth of
drawings and photos. In general, cladocerans (water
fleas) are a group of small animals belonging to one of
four crustacean orders (Table I). Adults are 0.2 –
18.0 mm long. The genera Daphnia (Fig. 1A) and Pleuroxus (Fig. 1B) exemplify typical cladoceran morphology. Cladocerans do not have a clearly segmented body
or appendages, although they do possess segmented
second antennae. Obvious features of most cladocerans
include a single central compound eye as adults and a
transparent clear-to-yellow carapace that is used as a
brood chamber. The carapace is attached to the back of
the neck and acts like an overcoat, wrapping around
851
part or all of the body, except for the head. In many
species, the 4 – 6 pairs of thoracic legs are covered by
carapace. Crustaceans differ from other arthropods in
having two pairs of antennae. In Cladocera, the first
pair of antennae (called antennules) are usually onesegmented and small, and have only a chemosensory
function. The second pair of antennae (called antennae)
are large and used for swimming.
Most cladocerans are small transparent animals,
whose general shape and jerky swimming accounts for
their common name, “water fleas.” Although abundant
in most standing freshwater, they are small enough to
be easily overlooked. Nevertheless, it is possible to observe their behavior and identify at least the larger
species using the unaided eye. For a clear view of their
structures, however, it is necessary to use a dissecting
microscope for gross features and a compound microscope for finer details.
Anatomy is best observed using living animals. At
first, it is easiest to focus on large and transparent animals, such as an adult Daphnia or Simocephalus. Once
structures are located in these large animals, they can
also be found in small animals, such as chydorids and
bosminids. Figure 1 indicates the appearance and relative arrangement of the major cladoceran structures.
For observation of a living cladoceran, one or
more individuals can be put in a small volume of water
in a Petri dish. If most of the water is then withdrawn,
the animals will be held down by the surface tension
and be unable to dart out of the microscope field of
view. For more careful observation, an animal can be
anchored to a small dab of clear grease. The grease can
either be put on the bottom of a Petri dish or on the tip
of a small glass rod.
Cladocerans typically are discus-shaped. When
dead or trapped by a surface film of water, they will lie
on their side. The body in side view (Fig. 1) is rounded,
the legs are obscured by a transparent shelllike covering (the carapace), and the major pigmented parts are
likely to be blue-to-yellow eggs in the brood chamber,
the green or yellow gut, and the dark black eye.
The location of the head is marked by one or more
eyes. The head is usually more or less dome-shaped but
can have long, even pointed extensions (compare
Fig. 2B and 2C) and an elongated beak may be present
(Fig. 1). All species possess a single black compound eye
except the chydorids Monospilus and Bryospilus
(Fig. 14 and 15) and the blind cave-dwelling species of
Alona and Spinalona (Dumont, 1995; Brancelj, 1990,
1992; Ciros-Perez and Gutierrez, 1997). The compound
eye is single, the result of fusion of two eyes during
embryonic development (Threlkeld, 1979). The black
color of the cladoceran eye is due to accessory screening pigments, including melanins, ommochromes,
FIGURE 1 (A) The anatomy of a daphniid. Daphnia pulicaria Forbes, 1893, emend. Hrbácek, 1959. Nebish
Lake, Vilas Co., Wisconsin, 6 June 1987. Figure drawn by Kandis Elliot (KE). (B) The whole-body anatomy of
a chydorid Pleuroxus trigonellus (O.F. Mueller, 1776). Rybinsk Reservoir, USSR. 7 December 1962. Redrawn
from Smirnov (1971) (KE); (C) Mandibular articulation in the chydorid subfamily Aloninae. Oxyurella brevicaudis Michael and Frey, 1983. D.G. Frey Sample 5019, Clearwater Lake, Putnam Co., Forida. 3 March 1979;
(D) Mandibular articulation in the chydorid subfamily Chydorinae. Pleuroxus aduncus (Jurine, 1820). D. G.
Frey sample 3004. Langemosa, Sealand, Denmark. 23 October 1972.
21. Cladocera and Other Branchiopoda
853
FIGURE 2 (A) Male Daphnia pulicaria Forbes, 1893, emend. Hrbáček, 1959. Nebish Lake, Vilas Co., WI, 6
June 1987; (B) Daphnia retrocurva Forbes, 1882. With elongated helmet. Lake Wingra, Dane Co., WI. 5
October 1978; (C) Ephippial female Daphnia pulicaria. Nebish Lake, Vilas Co. WI, 6 June 1997. (KE)
FIGURE 3 Male and female chydorid. Alona bicolor Frey, 1965. (A) Male redrawn from Frey (1965); (B)
female redrawn from Frey (1965).
pteridines, or purines (Shaw and Stowe, 1982). Since
the black color fades in several mounting media, such as
Hoyer’s, it is unlikely that much melanin is present. The
actual photoreceptive pigment is a nearly colorless purple, masked by the accessory pigments. With magnification of 50 , it is possible to see nearly transparent muscles that are attached to the eye and presumably
responsible for the motion. Once these muscles are seen
it is easier to find other muscles, as for example, those
attached between the basal segment of the swimming
(second) antennae and the back of the thorax and head.
Most cladocerans also have a small black ocellus (simple eye) just posterior to the compound eye. Like other
crustaceans, cladocerans show evidence of five pairs of
appendages on the head part of the body: two pairs of
antennae, one pair of mandibles, and two pairs of maxillae. At the base of the head, near the carapace margin,
is a pair of short cigarlike appendages — the first antennae (antennules). These are usually shorter than the
head and inconspicuous but may be longer than the
854
S. I. Dodson and D. G. Frey
head, as in Moina (Fig. 58) and most macrothricids
(Figs. 44 – 57). First antennae are not used for swimming or feeding but are probably chemosensory organs.
The head may curve toward and beyond the first antennae, forming a beak, as in Daphnia and chydorids
(Fig. 1), or the immobile first antennae can form a long
tusk, as in Bosmina (Fig. 42B). Attached to either side
of the body at the posterior margin of the head and
partly or entirely protruding from the carapace are the
second antennae (one on each side). These large
branched and segmented appendages are typically used
for swimming or crawling. The mandibles are unbranched and elongated, with a pivot point near the
carapace and a darkened grinding surface near the
mouth. The first pair of maxillae are typically reduced
to a small flap with a few curved spines. The second
pair of maxillae are missing or represented by a spine.
In addition to these appendages on the head, just above
the mouth is an upper lip (the labrum), which closes the
mouth and secretes mucus. A maxillary gland (nuchal
organ, or one or more head pores) opens on the dorsal
margin of the head in conchostracans and the cladocerans (Olesen, 1996). It secretes a glue in some forms
(Sida, Simocephalus), a large gelatinous capsule over the
carapace (Holopedium), or salt in marine or hypersaline
branchiopods.
The head smoothly joins the rest of the body which
is often covered by a carapace. The transparent carapace is often ornamented with a faint geometric pattern
in the planktonic species or by striae, spines, pits, or
hexagonal meshes in many of the littoral forms. The
cladoceran body posterior to the head is comprised of a
thorax and an abdomen hanging within the overcoatlike carapace. The body ends in a pair of (postabdominal) claws which can reach out of the carapace. Several
species have a stiff extension of the carapace: spines
near the base of the second antennae or teeth, spines,
or one long tail spine on the posterior end.
The chemistry of the carapace is important, since
cladocerans tend to have a hydrophobic exoskeleton.
Although surrounded by water, many cladocerans are
unwetted. If brought into contact with the surface film,
they become trapped. Their hydrophobic carapace is
probably an adaptation which discourages growth of
algae and protozoa on the carapace. Cladocerans cannot groom the outside of the carapace, and animals
that have lived for a few weeks without molting often
bear a thick load of encrusting organisms (Chiavelli
et al., 1993; Threlkeld and Willey, 1993) which may
diminish their feeding and swimming efficiency, reduce
their escape speed, and increase their sinking rate
(Allen et al. 1993).
Some cladocerans, such as Scapholeberis (Fig. 40)
and some forms of Daphnia, can be colored black over
parts of the body. The pigment is melanin, deposited in
the exoskeleton. The melanin is located in the parts of
the exoskeleton habitually faced toward the sun: the
margins of the carapace in Scapholeberis and the head
and back in Daphnia. Experiments have shown that
the black pigment provides protection against photodamage in Daphnia (Luecke and O’Brien, 1983), but
perhaps little photoprotection in Schapholeberis (Hurtubise and Havel, 1998). It is possible that induction of
melanin is associated with a decrease in reproductive
potential. It has also been proposed that the dark pigment also increases heating rate and, therefore, allows
cladocerans an increased reproductive rate in cold water. However, the small cladoceran body size and the
high thermal conductivity of water suggest that this effect is insignificant. Finally, while melanin strengthens
bird feathers and insect exoskeletons, it probably does
not have this effect in cladocerans (Dodson, 1984).
Melanistic animals brought into the lab lose their pigmentation at the first molt, suggesting that stimulation
of bright sunlight is needed to induce melanin.
Scapholeberis contain the darkest pigmented cladoceran species. Scapholeberis kingii is the only known
distasteful cladoceran; it is not even ingested by hydra
(Schwartz et al., 1983). Scapholeberis homogenate
causes Hydra to writhe and contract, and it induces
concerts of tentacular spasms. It is not yet known if
Scapholeberis is distasteful to visual predators. If so,
the black carapace pigmentation may be adaptive both
as protection against photodamage and as a warning
display.
The animal’s thorax and abdomen can be seen
moving within the carapace of living individuals. The
thorax holds four or six pairs of legs. These flat leafshaped legs (called “phyllopods”) are well supplied
with rows of setae and spines and are used for handling food, filtering, scraping, pumping, creeping, or
grasping females. The legs beat rhythmically back and
forth at roughly five beats per second at room temperature. If a small amount of an alga suspension is placed
with a pipette near the carapace margins, it is possible
to follow the water current which brings the algae to
the legs, and to watch algae collecting on the legs and
being passed toward the mouth. The thoracic legs
probably act as electrostatic filters (not sieves), collecting algae and other particles that stick to the flat surfaces and combs of setae (e.g., Gerritsen and Porter,
1982; Cheer and Koehl, 1987; Gerritsen et al., 1988).
Lampert (1994) showed that the filter comb size of
Daphnia (7 species) increased by as much as 83%
when the animals were grown at low food levels. Food
caught on the thoracic legs is mixed with mucus to
make a mass of food (the bolus), which is moved forward toward the mouth. The postabdominal claw is
21. Cladocera and Other Branchiopoda
used to clean the filters or to reject a bolus containing
distasteful algae. Cladocerans thus have the option of
accepting or rejecting a mass of algae, but little control
over the composition of the bolus (they have difficulty
making decisions about whether to accept each individual algal or bacterial cell, although Strickler developed
videos of Daphnia choosing individual particles). A bolus can be rejected if it contains a toxic alga or if its average “taste” is unacceptable (Richman and Dodson,
1983). If the food bolus is accepted, it is chewed by the
mandibles and swallowed. The amount of food ingested depends on the algal concentration, until a concentration is reached at which the legs are operating at
full capacity. Beyond this concentration, the amount of
food ingested remains constant, or even declines, as the
legs become clogged with food (Porter et al., 1982).
The mouth is near the margin separating the carapace and the head. On either side of the mouth lies a
pair of large but simple jaws (mandibles), each of
which is a single rod, pointed toward the outer end.
The inner dark chewing surfaces lie just in front of the
mouth. The jaws are in nearly constant grinding motion. If the animals have been feeding, the gut will be
easy to find. The gut runs from the mouth, loops
through the head, continues along the body through
the thorax and abdomen, and terminates at the anus
near the posterior end of the animal. The gut can be divided into three regions. The foregut and hindgut are
lined with cuticle, which is continuous with the cuticle
that covers the outside of the animal. The midgut (in
the thorax) is lined with epithelium elaborated into microvilli and is the site of absorption (Peters, 1987). In
the head region of Daphnia are a couple of small sacs
(hepatic caeca) attached to the gut. In some species, especially in the chydorids, there is a sac attached to the
gut in the abdominal region and none in the head region. A green to brown, rather amorphous material
typically fills the gut (Tappa, 1965) along with undigested algal cells and small animals (Porter, 1973,
1977). Waves of rhythmic (peristaltic) contraction
move along the gut from the anus toward the midgut.
By feeding a trapped animal two different kinds of
food (yeast followed by algae), it is possible to measure
the rate of movement of food particles through the gut.
Lying along the gut, in the thorax region of wellfed adults, are fat globules and a pair of gonads
(Fig. 1). The fat globules are light yellow to orange,
and are spread throughout the body cavity in greater or
lesser abundance (Tessier and Goulden, 1982). The
ovaries resemble a mass of large fat globules, but the
maturing eggs are more often slightly green in color
(the blue or green colors are due to carotenoids that are
combined chemically with proteins). Each time an
adult female molts, she extrudes a few new eggs into
855
the brood chamber, as she releases the neonates developed over the previous between-molt period. Developing sperm are large round cells, similar in appearance
to the fat globules. These are released through the
anus, when the postabdomen is inserted into the female’s brood chamber. When released, the spermatozoa
are amoeboid, without a flagellum. In most species they
are simple ovals 5 – 20 m in diameter. Spermatozoa of
Diaphanosoma are oval, about 60 m in diameter, and
those of Sida and Moina are spiny and nearly 100 m
long (Weismann, 1880; Wingstrand, 1978).
The heart (Fig. 1) is obvious because of its rapid
beating in live animals. It is a clear muscular organ lying in the thorax above the gut and just behind the
head. With the right light and at least 50 magnification, one can see the blood cells moving about, especially near the heart and in the bases of the second antennae, and in the beak of Daphnia.
The abdomen carries at its tip a pair of claws,
which may appear to be single. These claws are used to
groom the thoracic legs and can be seen to brush over
the thoracic legs as the abdomen is swung forward.
Some smaller species, especially those in the family
Chydoridae, use the abdomen as a sort of foot to kick
along surfaces.
The part of the body at the end of the abdomen
that bears claws and is posterior to the anus is called
the postabdomen. The anus lacks a muscle for closure
(sphincter). In live animals, rhythmic (peristaltic) waves
of contraction can be seen moving up the hindgut.
Most, if not all cladocerans show this “anal drinking.”
The hindgut cause a current of water to flow into the
anus; the water is absorbed by the columnar cells of the
anterior gut. This means that dissolved organic compounds may stay in the gut much longer than the food
from which they were derived (Fryer, 1970).
“Cyclomorphosis” (a type of phenotypic plasticity)
is a change in body shape within a species caused by alternate developmental pathways induced by environmental factors (such as temperature and turbulence)
and chemical communication (Dodson, 1989; Larsson
and Dodson, 1993). For example, shown in Fig. 2b is a
Daphnia with an elongated helmet. This same animal
could have offspring with much lower helmets, depending on the chemical signals present during embryogenesis. Animals from the same clone mother, but influenced by different chemical signals, have such different
forms that they have been described as different species
(Krueger and Dodson, 1981). The changes in form include elongated heads, tail spines, or in Bosmina, elongated first antennae. These elongated body parts are
sometimes accompanied by a reduced adult body size.
Black and Slobodkin (1987) define cyclomorphosis as
“temporal (seasonal or aseasonal) cyclic morphological
856
S. I. Dodson and D. G. Frey
changes that occur within a planktonic population.”
Animals generally lose elongation of body parts when
cultured in the laboratory for one or two molts. For example, Daphnia galeata mendotae and D. retrocurva
both produce higher helmets in warm turbulent laboratory environments. They also produce higher helmets
in the presence of two invertebrate predators,
Chaoborus larvae and Notonecta adults. Similarly,
Daphnia pulicaria produced neck teeth the first summer that Chaoborus were present in Lake Lenore,
Washington (Luecke and Litt, 1987). The induced morphological changes are generally thought to be advantageous for increasing survivorship in the face of specific predators: small size protects against fish and long
spines protect against small invertebrate predators
(Larsson and Dodson, 1993; Tollrian and Dodson,
1998).
2. Differences between the Sexes
The two sexes of cladocerans are morphologically
similar. Because most cladocerans reproduce asexually
at least part of the time, most individuals will be females. Males (Figs. 2A, 3B) resemble females but are
smaller. Cladoceran males have a copulatory hook on
the first thoracic leg, used for holding on to the female
during mating. Macrothricidae and Chydoridae have
the largest hooks, and Polyphemus and Leptodora
have the smallest hooks (Lilljeborg, 1901). Some cladoceran males have longer first antennae than are present
in the females (especially Sida, Diaphanosoma, Daphnia, Ceriodaphnia, Moina, Polyphemus, and Leptodora). Daphnia and Ceriodaphnia males have elongated setae on the anterior thoracic legs. Males of
Diaphanosoma, Latona, Bythothephes, Podon (Lilljeborg, 1901) and Penilia (Della Croce and Gaino, 1970)
have a pair of penes on the postabdomen, as do several
species of Alona and Leydigia.
3. Life History and Development
Detailed information exists for the life histories of
many cladocerans (Lynch, 1980; Frey; 1975, 1982c,
1987b, 1989) and especially for Daphnia (Schwartz,
1984; Lynch, 1989; Threlkeld, 1988). Cladoceran reproduce either sexually or asexually, depending on environmental conditions (Fig. 4). Most species reproduce parthenogenetically most of the time (Hebert,
1978, 1988), and most resting eggs develop into females. Species adapted to temporary habitats, such as
Moina species, have resting eggs that can develop into
either males or females.
Embryogenesis begins, and is usually completed, in
the brood chamber, a space between the body and the
carapace (Fig. 1, also see Kotov and Boikova, 1998). In
the Onychopoda and Haplopoda the carapace does not
FIGURE 4 Daphnia reproductive strategy. Adult females can produce
three different kinds of offspring, depending on environmental
conditions. Diploid (subitaneous) eggs are produced asexually, and
develop into females or males. Haploid eggs can also be produced,
given the correct chemical signal from the environment.
cover the body, but is reduced to a brood chamber on
the animal’s back (e.g., Figs. 60 and 61). Cladoceran
eggs (Figs. 1, 7, 9B, 13, 27, 56, and 61) are of three
types:
• diploid “subitaneous” eggs, which develop immediately into young;
• resting “eggs,” which come from haploid eggs
that are fertilized, develop into early embryos,
and then enter a diapause (state of physiological
stasis) which is resistant to heating, drying, and
freezing; and
• pseudosexual resting eggs, which are early
diapausing embryos from asexually produced
diploid eggs (no fertilization needed).
Why do cladoceran species often reproduce both
asexually and sexually? Two adaptive advantages of
asexual reproduction are that population growth rate is
maximized, since all offspring are females, and genetic
combinations particularly well adapted to present conditions are not disrupted by sexual recombination
(Banta, 1939; Williams, 1975). Two disadvantages to
asexual reproduction are that asexual clones are slow
or unable to adjust to changing conditions by sexual recombination (and so are restricted to specific habitat
conditions) and that asexual genotypes accumulate
mutations (which are typically detrimental). Sexually
21. Cladocera and Other Branchiopoda
derived resting eggs represent new genetic combinations
and have a chance of both being pre-adapted to new
environmental conditions and lacking detrimental
mutations.
The different adaptive advantages of the sexual
and asexual eggs would seem to dictate that only the
sexual eggs should diapause, as is often the case. The
exception to this is the production of “pseudosexual”
resting eggs in obligate asexual clones. Pseudosexual
resting eggs are produced without recombination but
otherwise resemble sexual diapausing eggs and are enclosed in an ephippium. Pseudosexual eggs are probably a solution for the need to produce resting eggs in
ponds (such as in the arctic or alpine zones) that are
too cold to allow enough time for two generations of
Daphnia (one generation of females out of the resting
eggs, which produce males and females, which in turn
produce new resting eggs). Pseudosexual eggs can be
produced by the females that emerge from resting eggs.
“Asexual, immediate reproduction” is achieved via
subitaneous eggs, which have a visible yolk center and
are often pigmented yellow, blue, or green. Subitaneous
eggs develop into neonates inside the mother’s brood
chamber. Egg developmental stages can be followed
easily because the carapace is transparent (Threlkeld,
1979; Kotov and Boikova, 1998). The neonates are released from the brood chamber as the mother molts.
After molting, she may extrude another set of eggs into
the brood chamber, depending on her age, nutritional
status, and the environmental conditions.
Except for Eurycercus and two other genera, the
chydorids have a constant clutch size of two; whereas
in other cladocera, clutch size increases with body size,
so that the smallest species produce one or two eggs
per clutch while the largest species can produce hundreds of eggs per clutch (Hann, 1985). Small individuals carry only a few eggs (which are huge relative to the
adult, as in Fig. 1D). Reproduction is very expensive
for small cladocerans. The small Daphnia lumholtzi
typically carry two eggs in the first reproductive instar.
These eggs account for 30 – 80% of the female’s weight
(King and Greenwood, 1992). Large Daphnia individuals (such as Daphnia magna) also allocate a significant
proportion of their energy intake to reproduction; but
because they are larger and produce relatively smaller
eggs (not much larger than those of the small species),
they can carry dozens or even hundreds of subitaneous
eggs. These eggs begin developing as soon as they are
extruded into the brood chamber. Egg development
time and longevity are proportional to the inverse of
temperature to an exponent of about 2.5 (Rigler and
Downing, 1984).
“Delayed reproduction” is often associated with
sexual reproduction (Hebert, 1978, 1988). Males are
857
diploid and produced asexually, just as their sisters are.
Production of males is induced by some environmental
signal, such as a change in food concentration, a chemical signal produced by crowded Daphnia, or changing
(usually decreasing) photoperiod (Ferrari and Hebert,
1982; Hobaek and Larsson, 1990; Kleiven et al., 1992,
see Fig. 4). When a population begins reproducing sexually, females produce mixed clutches of males and females. The male to female ratio varies greatly between
populations and in different years. Highest sex ratios
are typically seen in small ponds, and there is some
concern that environmental contaminants may have reduced male production (Dodson and Hanazato, 1995).
Sexual reproduction begins when one or two haploid eggs are extruded into a modified (thickened) carapace (except in the Sididae and the predaceous species)
called the ephippium (in Greek “saddle”, see Fig. 2C).
The changes in carapace development are induced by
either environmental (chemical) signals from crowding
or toxic food (Ferrari and Hebert, 1982; Carvalho and
Hughes, 1983; Shei et al., 1988; Kleiven et al., 1992)
or by environmental contamination (Van Der Hoeven,
1990; Shurin and Dodson, 1997). Ephippia are produced most often in temporary ponds, especially in
harsh environments (Lynch, 1983; Weider and Hebert,
1987).
Upon fertilization of the haploid egg by the male
(Jurine, 1820; Baird, 1850), the resulting zygote (or the
diploid pseudosexual egg) undergoes several cell divisions (Banta, 1939; Ojima, 1958) and then enters diapause. These resting “eggs” (actually early embryos)
are resistant to freezing and drying and are further
protected by the thickened part of the carapace. The
ephippium is darkly pigmented with melanin. The
rough surface of the carapace may serve as a speciesspecific recognition cue for the male during mating.
The ephippium is shed when the female molts. The
ephippium closes around the resting eggs, and most of
the rest of the exoskeleton detaches from the ephippium (except for the tail spine in Daphnia).
In some populations, resting eggs are not the result
of sexual reproduction but are produced asexually
(Hebert, 1987; Weider and Hebert, 1987; Innes and
Hebert, 1988). These “pseudosexual” resting eggs are
not produced by sexual reproduction, but are the result
of a gene that suppresses meiosis. They act just like the
sexually derived resting eggs, but are genetically identical to their mother.
Chydorid and macrothricid ephippia sink and are
usually attached to the substrate (Bretschko, 1969;
Fryer, 1972). Daphnia ephippia tend to float, because
of their specific gravity and the hydrophobic character
of the carapace. Dark pigmentation of ephippia may
aid dispersal, since fish, amphibian, and bird predators
858
S. I. Dodson and D. G. Frey
can easily see the dark ephippium and the ephippium is
not damaged by passage through a predator’s gut
(Mellors, 1975). The tail spine and, sometimes, a strip
of chitin ventrally at the head end of the carapace
(Fryer, 1972) may aid dispersal by allowing the ephippium to stick to fur or feathers of wading animals.
Ephippia are small enough to blow about with the
wind and resistant enough to remain alive on dry land
for years (Moghraby, 1977).
“Egg banks” are composed of resting eggs stored
in the sediments or watershed. The resting eggs may be
years or decades old, and they appear to hatch leisurely
over several years. Egg banks provide an important
large-scale stabilizing influence on the genetic structure
of a cladoceran population and a hedge against extirpation (Hairston, 1996). To continue development, the
diapausing embryos in ephippia require stimuli such as
water, long or increasing day lengths, and high oxygen
concentrations (Schwartz and Hebert, 1987a). The
resting eggs usually develop into females which produce more young asexually. Major exceptions are
Daphniopsis and Moina, genera which specialize on
short-lived temporary ponds and which produce both
males and females from the ephippial eggs (Goulden,
1968; Schwartz and Hebert, 1987b).
Molting allows cladocerans to increase in size. As
the embryos develop, they assume the appearance of
the adult. After release from the brood chamber, the
newborn (neonates) can grow only by molting. That is,
when sufficient energy has been stored, the animals: (a)
reabsorb the inner part of their exoskeleton; (b) begin
to manufacture a new exoskeleton below the old; (c)
absorb water; and (d) pull themselves out of the remaining outer layers of the old exoskeleton, exiting at
the back of the neck. The old exoskeleton (the exuvia)
is a transparent ghost image that retains the complexities of the outer surface of the animal, including all the
fine setae of the thoracic legs. Exuviae have been used
to measure changes in size of various structures as an
animal grows from neonate to adult. The energetic cost
of molting may be a major constraint on body size in
Daphnia (Lynch, 1989). Given a limited rate of food
intake, cladocerans may be faced with the choice of allocating energy to molting (to become larger) or allocating energy to reproduction.
The developmental stage between molts is called an
instar. There is some variation in (and controversy
about) how many times cladocerans molt. Neonates
molt just twice in chydorids but up to seven times in
daphnids before producing eggs of their own. The number of juvenile instars is more or less species-specific and
depends on adult body size, with the smaller species
having the smallest number of molts. The rate of molting (instar duration) in Daphnia pulex depends mostly
on the temperature, less on food (Lynch, 1989). If animals are poorly fed, they may even lose weight when
they molt. An instar lasts from about a day to weeks,
depending on the temperature. In laboratory studies,
cladocerans can live for months at room temperature.
Each time a well-fed juvenile molts, it approximately doubles its volume, and its length increases
roughly by a factor of 1.26. Growth slows as cladocerans mature; successive adult instars grow slightly, but
energy not used for maintenance is allocated to produce eggs instead of growth (Frey and Hann, 1985).
Female cladocerans can molt several times after becoming adults. Chydorid males become mature in the third
instar and stop molting, although Daphnia males continue to molt and grow after maturity.
4. Induced Changes in Life History
Many cladoceran life-history characteristics include instar duration, number of instars to first reproduction, time to first reproduction, primiparous and
neonate size, and clutch size. These characters have a
genetic basis (Spitze, 1995) and show trade offs and
correlations that are suggestive of adaptations to the
environment and to biological interactions, especially
predation.
Life-history characteristics also show phenotypic
plasticity (Hann, 1984; Schwartz, 1984; Larsson and
Dodson, 1993). That is, the same genotype can develop
differently, as it is influenced by temperature, food supply, and chemical signals produced by predators. The
patterns of phenotypic response (reaction norms) appear to be, in general, adaptive responses to size-selective predation and food limitation.
B. Metabolism
The many physiological studies of Daphnia provide
a reasonably complete picture of cladoceran metabolic
rates (e.g., Peters, 1987). Metabolic rate is measured as
the rate at which an animal uses oxygen or produces
carbon dioxide. As long as oxygen is present, gas exchange (oxygen in and carbon dioxide out) is not a
problem for small animals, because of their high surface-to-volume ratio. Gas exchange probably takes
place across the entire surface of the animal, not just on
the gill like thoracic legs. Porter et al. (1982) and Peters
(1987) found no correlation between rate of leg movements and respiration rate. Animals collected in oxygen-poor habitats (sewage lagoons or temporary pools
with decaying vegetation) will often have a general
pink-to-red color (Weider and Lampert, 1985). This is
due to hemoglobin dissolved in the blood. If a red animal is wounded, it will bleed red blood; and if the
blood is examined at high magnification, the blood cells
21. Cladocera and Other Branchiopoda
859
will appear less pigmented than the surrounding fluid.
Animals become clear in a day or so if grown in welloxygenated water, suggesting an energetic cost to maintenance of hemoglobin (Landon and Stasiak, 1983).
Metabolic rate is influenced by a number of factors, including temperature, body size, and food concentration.
can be ingested. In the absence of food, Daphnia
continue to move their legs, as if the thin gill-like legs
were also being used as respiratory organs, requiring
ventilation.
1. Temperature
Environmental temperature may be a causal factor
for body size. For example, Stirling and McQueen
(1986) found that the direction of change in environmental temperature was correlated with body size and
reproductive strategy in Daphniopsis ephereralis, an inhabitant of cool forest ponds. Annual changes in average body size for the population was not directly related
to predators and may have a physiological explanation.
Starvation is a problem faced by most cladoceran
populations sometime during each year (Threlkeld,
1976, 1988). Because smaller animals have a higher
metabolic rate per unit body weight, neonates are the
more prone to starvation than adults, if no food is present (Tessier and Goulden, 1987). Eggs can receive a
large store of energy-rich fat from the mother, if the
mother is well fed. However, this energy store is reduced if the adult is faced with limited food. Similarly,
if food is absent, the large species can persist for longer
than small species. Even for large adults, a single day
without food results in a drop in energy stores, body
weight, and respiration rate.
At food concentrations above starvation levels, the
relationship between food availability, energy allocation, and body size are complex. How do cladocerans
allocate the energy gained from limited food input?
Models of energy allocation based on physiological
principles suggest that energy is used for different purposes at different life stages (for example, only for
maintenance and growth in immature animals), and
that adults have allocation priorities for maintenance,
reproduction, and growth (Bradley et al., 1991).
Large animals always have lower weight-specific
respiration rates. However, smaller animals have the
higher weight-specific assimilation at low food concentrations, and large animals have higher rates at high
food concentrations (Tessier and Goulden, 1987). This
suggests that net growth, estimated by the difference between assimilation rate and respiration rate, is highest
for small animals at low food concentrations and highest for large animals at high food concentrations. In a
study of three Daphnia species (Tillmann and Lampert,
1984), the large D. magna out-reproduced smaller
Daphnia. When food was marginally limiting, the three
species had similar reproductive rates; and when food
was severely limiting, only the smallest species, D.
longispina could still reproduce. Enserink et al. (1996)
also found that larger individuals appeared to grow
Metabolic rate has a hump-shaped (inverted-U) relationship with temperature. At some low threshold
temperature, metabolic rate is just fast enough to maintain growth and reproduction. A rise in temperature increases metabolic rate up to a maximum. Above the
temperature of maximum metabolic rate, further increases cause death, either because critical metabolic
enzymes are thermally damaged or because respiration
rate exceeds the rate of energy intake (Moore et al.,
1996). Most cladocerans are unable to live at temperatures above 30°C, and animals native to cold waters
may have much lower upper thermal limits. This sensitivity to high temperature can be increased if the cladocerans are feeding on toxic bluegreen algae (Threlkeld,
1986a). Both Ceriodaphnia and Diaphanosoma live
naturally in water of 27 – 30°C. When bluegreen algae
are present, Ceriodaphnia shows a decline in population growth rate. For short periods of time (15 min),
Daphnia can tolerate temperatures up to about
35 – 38°C, depending on the temperature at which they
were raised (MacIsaac et al., 1985).
2. Body Size
Small animals, cladocerans have relatively fast
rates of metabolism. Richman (1958) found that respiration rate of Daphnia is proportional to body length
raised to the 2.14th power, a value similar to size-specific respiration rates reported for many invertebrates
(Schmidt-Nielsen, 1984). The range of cladoceran body
size is enough to ensure significant differences in metabolic rates between the smallest and largest cladocerans. While larger animals require more oxygen and energy in an absolute sense, large animals use less energy
and oxygen per unit of weight.
3. Food Concentration
As food becomes more concentrated, the animals
respire at an increasing rate, probably because of increased metabolic needs: assimilation rate also increases
with food concentration (Peters, 1984; Lampert,
1986a). When food is superabundant, the grooming required to clean the filters on the thoracic legs may also
increase the respiration rate (Porter et al., 1982). This
phenomenon can lead to starvation at concentrations of
food high enough so that the increased respiration requirements exceed the maximum rate at which energy
4. Interaction of Temperature, Body Size,
and Food Concentration
860
S. I. Dodson and D. G. Frey
fastest at constant low food levels, whereas smaller individuals had the advantage with fluctuating food.
When a cladoceran is food limited, it tends to mature at a smaller size and produce smaller (but as
many) offspring. The main response of Daphnia pulex
to low food levels was a reduction in size-specific food
intake and egg size (Lynch, 1989). Over a wide range,
food concentration had no effect on length-weight relationship, instar duration, or weight-specific investment
of energy in reproduction.
Population growth rate is linked to body growth
rate. Competition among various species of cladocerans may largely depend on which species can grow
fastest at a given food level. Thus, small species such as
Ceriodaphnia and Bosmina may be able to out-compete, or at least out-grow, large species such as Daphnia pulex at low but not at high food concentrations
(Tessier and Goulden, 1987).
III. DISTRIBUTION
A. Zoogeography
Cladocerans are a widespread group occurring in
all but the most extreme freshwater habitats (Hutchinson, 1967). While they are more abundant in lakes,
ponds, and slow-moving streams and rivers (Thorp
et al., 1994), they also occur in quiet water and marginal vegetation in fast-moving streams. Several species
probably occur in groundwater, especially river gravels
(Dumont, 1987). One chydorid cladoceran has been
found in water trapped in mosses and bromeliads that
drape trees in the cloud forest of El Yunque, Puerto
Rico (Frey, 1980b). One species or another are found
from sea level to ponds above tree line, and from arctic
to tropical latitudes. In North America, the highest
diversity of planktonic species is found in the glaciated
midtemperate zone (Tappa, 1965), and the highest
diversity of benthic species, especially small-bodied
species, is found in the southeastern United States
(Crisman, 1980; Kerfoot and Lynch, 1987).
Chydoridae, now seems to be mainly an illusory concept (Frey, 1987a), based on an incomplete appreciation of the morphology of seemingly the same taxa in
different places and on an unquestioning acceptance of
the efficiency of passive dispersal of these organisms via
their resting eggs as carried by wind, water, birds, insects, and mammals (Frey, 1982a). Recent studies in
North America (Frey, 1986a) are demonstrating that
the geographical ranges of species of chydorids can be
just as narrow and circumscribed as those of animals
considered to be much less vagile. Careful genetic
analysis (Hebert, 1987) has shown Daphnia species to
be more diverse and to have distributions restricted to
small portions of North America. In fact, those Daphnia species that are obligate parthenogens are genetically very diverse (Hebert, 1987), nearly as diverse as
asexual ostracods (Havel and Hebert, 1989; Chaplin,
1994). Thus, the cladocera of different continents are
probably different species (even though many now
have the same specific names), and the distributions of
species within a continent depend on the distribution of
favorable environmental conditions.
Hybrids between cladoceran species are often reported (Hebert, 1985; Hann and Hebert, 1986;
Hebert, 1987; Mort and Streit, 1992; Taylor and
Hebert, 1993; Spaak, 1995; Colbourne et al., 1997).
The evidence has until recently been only morphological, or based on lab crosses. For example, Pleuroxus
denticulatus and P. procurvus were crossed in the laboratory to produce one individual which could only reproduce asexually and was morphologically intermediate between its two parents (Shan and Frey, 1983).
Recent studies based on molecular characters confirm
that hybrids can indeed be formed between similar
species. The persistence of clones in nature is due to
their ability to reproduce asexually. Species remain distinct, in part, because the hybrids are not known to be
able to reproduce sexually in nature. Thus, while
cladoceran species are often difficult to distinguish using morphological characters, molecular characters
show them to be clearly distinct.
C. Biodiversity
B. Cladocera and the Species Concept
Our understanding of the distribution of cladoceran species depends on how well we can distinguish
species. Cladoceran taxonomy is currently undergoing
a reanalysis. Because of an almost complete disregard
of fine but significant details of morphology and a consequent impression that taxa on different continents
were the same, many investigators in the past (Forbes,
1925) and present have claimed that many species are
cosmopolitan. But cosmopolitanism, at least among the
One measure of biodiversity is the number of
species, also called “species richness.” Cladoceran
species richness in a lake depends on several factors, including water chemistry, lake size, productivity, the
number of adjacent lakes, and biological interactions
(Dodson et al., 1999).
1. Water Chemistry
Carter et al. (1980) found that of 23 pelagic cladoceran species occurring in eastern Canada, eight may
21. Cladocera and Other Branchiopoda
have distributions restricted by water chemistry: Ceriodaphnia and a species of Eubosmina tended to occur in
hard water, and Polyphemus, Sida, two Daphnia
species, and two other Eubosmina species were found
mostly in soft water.
Salinity is a major factor that limits cladoceran distribution. Cladocerans as a group have a wide range of
tolerance to salinity (Potts and Durning, 1980), but individual species are adapted to a relatively narrow
range of salinity (Teschner, 1995). Particular species can
be found from near-distilled water (Dodson, 1982) to
the 3.2% salinity of oceans (Della Croce and Angelino,
1987), to water more saline than sea water. For example, Moina lived in Soap Lake Washington at salinities
as high as 39 g / L (Edmondson, 1963). High salinity excludes most cladocerans while providing a refuge for
physiologically specialized genera such as some Moina.
The cladocerans of Pyramid Lake, Nevada, showed
strong mortalities at salinities of 0.6% (Diaphanosoma), 0.7% (Ceriodaphnia), and 1.8% (Moina)
(Galat and Robinson, 1983). The marine cladocerans
Evadne and Penilia are restricted to coastal and open
waters, while Podon is found in less saline coastal and
estuarine water (Della Croce and Angelino, 1987).
Even within a species, there is evidence of adaptation to average salt conditions. For example, different
clones of Daphnia magna, from fresh or brackish ponds,
show differences in their level of salt tolerance and do
poorly when raised at different salinity levels than their
native pond (Weider and Hebert, 1987; Teschner, 1995).
Acidity also affects cladoceran distribution. Cladocerans are mostly found in neutral or alkaline water,
although a few live in acidified lakes. Only a few
small cladocerans such as Bosmina, Chydorus, Diaphanosoma, small Daphnia, Holopedium, and
Polyphemus can tolerate extremely acidic water, occurring in lakes with pH values between 5.0 and 3.8
(Sprules, 1975; Frey, 1982c).
Turbidity affects cladoceran distribution, probably
because clay particles interfere with cladoceran filters
(Gerritsen and Porter, 1982). A combination of laboratory life table experiments and field observations suggested that Moina and Diaphanosoma are adapted to
silt-laden water, while Ceriodaphnia and Daphnia
(subgenus Daphnia) are not (Threlkeld, 1986b). Some
species of Daphnia (Ctenodaphnia) (Dodson, 1985)
and several chydorids (such as Graptoleberis) are
found in very turbid water or in mud.
2. Lake Size, Productivity, and Adjacent Lakes
For pelagic cladoceran species, there are more
species in larger lakes, with about 35 species in the
largest freshwater lakes. Small ponds and pools (as
small as a few liters) may have one or two species. As
861
lake size increases, there are probably more distinct
habitats for different species. The highest species richness occurs in mesotrophic lakes with average annual
primary productivities of about 100 g C / m2 per year.
The mechanism by which productivity affects cladoceran species richness is still unclear. Lakes surrounded
by lakes have a higher species richness than isolated
lakes (such as Crater Lake in Oregon). Clustered lakes
probably act as sources for each other for immigrants
(usually via resting eggs) among lakes.
Whether a species occurs in a body of water depends partly on the: (a) zoogeographic region, that is
on the probability of getting into the water; (b) chemical and physical requirements of the species; (c) food
conditions within the water; and (d) predators living in
the water. Although it is believed that some cladocerans
are capable of long distance dispersal via resting eggs
carried by the wind and by feathers and guts of water
fowl, we still know little about actual rates of colonization. Inbreeding coefficients for small Daphniopsis
populations suggest that populations receive an average
of 0.3 migrants per generation (Schwartz and Hebert,
1987b). Studies of Daphnia indicate slightly higher
rates of migration of about 0.5 migrants per generation. However, studies of several arctic cladocerans
suggested that dispersal rates are quite low, and that
pond populations, even when near each other, retain
genetic differences (Boileau et al., 1992).
3. Biological Interactions
Cladocerans compete for food with other cladocerans and with other herbivores (Threlkeld, 1988). In
some cases, this competition is intense enough to result
in marked reduction in abundance of other cladocerans
and potentially to result in competitive exclusion.
Vanni (1986) found that Daphnia could reduce algae
concentrations to a level that caused Bosmina to decrease. He suggested competition for food is a cause of
the often observed scarcity of small species in lakes inhabited by large cladocerans such as Daphnia. This effect may be especially strong in nutrient-rich lakes,
where Daphnia populations can increase rapidly
enough to produce a sudden and major reduction in algal abundance.
Size selective predation sometimes affects the distribution of cladocerans (Zaret, 1980; Kerfoot and Sih,
1987). In general, large-bodied species tend not to coexist with fish and small-bodied species can be excluded by small predators such as the larvae of the
midge Chaoborus or the backswimmer Notonecta.
Thorp (1986) analyzed 18 reports of predator – prey
community experiments in freshwater zooplankton. He
found that adding predators often reduced the number
of individuals of cladocerans present, but that there are
862
S. I. Dodson and D. G. Frey
few studies showing an effect on the number or relative
abundance of prey species. Over the long term, predators may influence cladoceran diversity via coevolution,
size-selective predation, and restrictions on prey distributions.
While food limitation and selective predation are
components determining the distribution of zooplankton, it is clear that the two factors are intertwined, that
predators influence competition and vice versa via what
are coming to be called “indirect effects” (Cooper and
Smith, 1982; Lampert, 1987; Kerfoot and Sih, 1987;
and Carpenter, 1988). Fryer (1985) concluded that diversity of chydorids within a lake depends only slightly
on dispersal, competition or predation, and more on
microhabitat preferences and presence of the appropriate specific food.
4. Littoral and Meiobenthic Species
We know much more about pelagic species richness
and distribution than we know about the littoral (or
meiobenthic) species, such as the chydorid, macrothricid, and sidid cladocerans. The meiobenthic cladocerans
comprise a major part of the diverse littoral community
of lakes, which includes such additional metazoans as
cyclopoid and harpacticoid copepods, occasionally
calanoids as well, ostracods, amphipods, rhizopods, hydra, bryozoans, rotifers, mites, nematodes, oligochaetes,
and larval or nymphal stages of midges, corixids, notonectids, beetles, caddisflies, mayflies, damselflies, as
well as some other groups. Besides this high diversity of
animals, there are the various macrophytes, sessile and
substrate-associated algae, and bacteria. Of all the
groups of animals, the cladocerans typically are most
abundant, at times reaching densities greater than 1 million per square meter of bottom, even under the ice in
early winter. The composition of this littoral community
and its importance relative to the offshore planktonic
and benthic associations varies with the geographic location of the lake and its general characteristics of size,
depth, and transparency.
There are typically about 8 – 15 species of meiobenthic cladocerans present in a given habitat, sometimes
five or fewer and occasionally 25 – 30 or even more. As
with pelagic cladoceran species, the species richness of
littoral cladoceran species appears to be correlated with
lake size. For example, in a study of 207 water bodies
in Yorkshire, England, from small ponds (5000 m2)
to small lakes (15,000 m2), Fryer (1985) found more
littoral cladoceran species in the larger lakes. Small
ponds had a median of about eight species; the larger
lakes had about 30 species.
Some littoral species are associated only with the
sediments and others with macrophytes. Some species
occur only in acid, softwater habitats, others in quite
strongly alkaline ones. Fryer (1968) showed that the
species he studied are diversified as to food eaten and
how this is obtained. One species of chydorid is a scavenger, another scrapes food from the bottom as a snail
does, and the third an ectoparasite on hydra. Most of
the remaining species use the same common food supply on substrates — organic detritus with its contained
algae and bacteria. The structure of the five pairs of
trunk limbs is remarkably uniform among all these
species, suggesting little variation in method of getting
the food. Most specializations within chydorids seem
linked to obtaining food from slightly different microhabitats.
Meiobenthic cladocerans occur virtually wherever
water is present, from water bodies of “normal” size
down to marshes, swamps, pools, puddles, ditches, seasonal water bodies, groundwater, running water of all
sizes and nearly all velocities, moist sphagnum, and
even in overgrowths and cushions of leafy hepatics in
rain and cloud forests, up to several meters above the
forest floor. Most species are sensitive to salinity, although a few occur at concentrations up to 5 – 10%
(g / L), sometimes even higher.
5. Conservation Issues
Unlike the large branchiopods (see below), conservation issues do not exist for cladocerans. Rare species
probably exist, as do opportunities to conserve cladoceran biodiversity. However, rare cladoceran species
are probably morphologically similar (or identical) to
common closely related (sibling) species. Only a combination of morphological and molecular analysis is
likely to identify rare cladoceran species (for an example of a fruitful protocol, see DeMelo and Hebert,
1994, and Hebert and Finston, 1996).
As in other areas of conservation, the invasion of
exotic species is a concern for North American communities. For example, Daphnia lumholtzi was probably
introduced into North America from Africa, along with
Nile perch. It is now a major component of lake and
reservoir plankton communities in southern North
America (Havel, 1995) and occurs in some rivers, including the Ohio (Jack and Thorp, 1995). Besides being introduced along with fish, cladocerans invasions
of new continents are either mysterious (Benzie and
Hodges, 1996) or have been connected to heavy (military) equipment (Schrimpf and Steinberg, 1982).
IV. BEHAVIOR
Cladocerans display a number of behaviors relating to mating, escape from predators, feeding, and
movements within lakes.
21. Cladocera and Other Branchiopoda
A. Mating Behavior
Jurine (1820) provides us with one of the few descriptions of mating behavior in cladocera. In November 1797, in the region of Geneva, he found a pond
containing a dense population of Daphnia, which included ardent males. He observed numerous couplings
in the laboratory, and reported (translated from the
original French):
“The male jumps onto the back of the female, which sometimes
escapes, but if he is able to seize her with the long filaments of
his first legs and secure her with his elongated first antennae, he
is able to catch hold of her firmly. He soon scuttles quickly over
the surface of her carapace until he has attained the lower
margin. When he finds the position so that the openings of the
two carapaces are opposed, he quickly introduces into her
carapace the long setae of the first antennae and of the first
thoracic legs, and thereby envelopes and binds, so to speak,
the legs of the female. When he is established in this position,
he curves his postabdomen forward, putting it out far enough
to get that of the female. When she feels that part, she is greatly
agitated, and fleeing, carries the male so fast it is hard to follow
the amorous couple in the vase that contains them. Finally, this
agitation ceases and the female in turn advances her
postabdomen in order to touch that of the male. The
postabdomens hardly touch before they separate. At the instant
of this touch, the male is agitated by spasms which give his legs
remarkable vibrations. It is during this contact that, in my
opinion, the copulation occurs.
The embrace is of variable duration; it rarely is sustained for
more than eight or ten minutes. During this interval, the
postabdomens were brought together more than once.
Copulation terminated, the postabdomen of the male is slowly
withdrawn into his carapace, but he is not yet able to withdraw
the setae of his first antennae and first thoracic legs because of
the spasms that persist in these parts. When these cease, the
separation of the two animals takes place.
This is not the only mode of embrace in these animals. I have
several times seen the male introduce only one first antenna, one
first leg seta, and one swimming (second) antenna with which
he envelopes the setae of the females second and third legs.
When the copulation is finished, he is not able to retrieve his
swimming antenna before the spasms which he is experiencing
are entirely gone.
I noticed other animals that were embraced almost transversely,
in such a way as to form a sort of cross.
Sometimes other males arrived intending to unite with a female
already in an embrace. One then sees them explore the female’s
body with a remarkable vivacity, trying, although futilely, to
penetrate into the sanctuary of pleasure, by elongating their
postabdomen as much as they can into the opening of the
carapace.
In all cases where copulation had taken place, the postabdomen of
the male returned slowly and with shudders to its usual position,
frequently interrupted by convulsive movements which stretched
it out to re-approach the female, such that it took a minute or two
for the postabdomen to be completely at rest again.
Males attacked indiscriminately all females they encountered.
Despite that, I assume that true copulation took place only with
those females that had no eggs in the brood chamber, for with
these the act was reiterated and prolonged. For the others,
863
although the preliminaries were the same, they terminated
very quickly. Ephippial females pushed aside the males and
rendered their attacks in vain.”
About 60 years later, Weismann (1880) gave cursory descriptions of mating behavior for Moina, Daphnia, and Bythotrephes, and conjectures on possible
mating behavior, based on anatomy, for Daphnella, Latona, Sida, Holopedium, Ceriodaphnia, Scapholeberis,
Simocephalus, Bosmina, Pasithea, Macrothrix, Eurycercus, Pericantha (now Pleuroxus) truncata, Chydorus,
Podon, Evadne, and Leptodora. These mid-Victorian
descriptions lack the enthusiastic attention to behavioral details shown by the pre-Victorian Jurine (1820),
but do include a wealth of detail concerning cellular
and tissue structure and ontogeny
“Mating in Daphnia has been observed by many, although only
the external association of the genders: the attachment of the
male and the combined swimming about. Indeed, Jurine and
after him many others noted that for Daphnia pulex, the ardent
(begattungslustigen) males attacked nearly every female,
whether they are carrying ephippia or embryos. This is indeed
correct as far as it goes, but they always quickly let them go
again and find a true mate — as far as I can judge — only
succeeding with a female that carries a winter (haploid) egg in
the oviduct. Sometimes the males are not so ardent, and wait
for the correct moment. Thus, on the second of September
1875, I brought several Daphnia pulex males together with a
female which carried ripe embryos in the brood chamber and a
newly ripe haploid egg in each ovary. The males made no
advances during the entire morning. First about 4:30 pm, after
the female had delivered her young, a male attached himself on
the back of the carapace, and quickly within a few seconds,
hitched himself around in front. The female did not defend
herself, but swam peacefully and slowly about, also sometimes
lying still on the bottom. The male did not let go for more than
a quarter of an hour. Sadly, I had to interrupt the observations,
and when I was able to take them up an hour later, the mating
was already consummated, the male had departed, and in the
brood chamber, which had not yet developed an ephippium, lay
two fertilized winter eggs . . . .” Weismann (1880) further
argued that successful fertilization can occur in Daphnia only
with the partners face to face and parallel.
The veil of modesty is further drawn over mating
behavior by monographers of functional morphology,
species, and evolution who largely ignored the topic of
mating behavior (e.g., Banta, 1939; Brooks, 1957;
Fryer, 1968; Goulden, 1968; Fryer, 1974). However,
perhaps influenced by the renaissance of natural history
in the last couple of decades and the general freeing-up
of sexual attitudes of the late sixties, Shan (1969) and
Smirnov (1971) have produced much more informative
descriptions of chydorid mating behavior and functional morphology. The theme of female participation,
and even perhaps female choice, introduced by Jurine
(1820), is developed further by Shan (1969):
“My observations on laboratory stocks of P. denticulatus do
not agree with Weismann’s (1880) description. At the
864
S. I. Dodson and D. G. Frey
beginning, the male swims vigorously here and there, using his
first antennae, trying to find a female that is receptive, usually
an ephippial female. A receptive female often cooperates in the
copulation process. Sometimes, the ephippial female he
approaches may not be receptive and she may reject him using
her postabdomen as he approaches her. At times a male may
hook onto a parthenogenetic female and try to copulate with
her, or a male may hook onto another male. In one occasion, I
saw a parthenogenetic female that had three males attached in a
series, one after another!
As the male hooks onto the edge of the female’s carapace, he
moves vigorously back and forth along the edge, often only on
one side of the female. The contact points are the two
copulatory hooks [on the first thoracic leg] and the tip of his
rostrum. He jerks his postabdomen and moves his hooks
vigorously while the female does not react to him in any way.
This activity goes on for a while, usually a few minutes,
possibly even longer than ten minutes. The female may become
excited when the male reaches a position where his hooks are at
the top of the free posterior edge of the female’s carapace. In
this position the male and the female are facing the same
direction and have the same dorsal orientation. At this time the
female may stop feeding, carry the male with her, and swim
away. They swim together, both jerk their postabdomens and
strike their antennae. They swim for some time, possibly ten
minutes or longer. Finally, they fall onto the bottom, still
hooked together, and lie on one side. Both of them stretch and
jerk their postabdomens back and forth. At this time the
female’s postabdomen is in maximum extension downward and
backward, while the male is flexing his forward trying to get as
close as possible to the brood pouch of the female. Presumably
at this time the sperm are ejected. The mechanism that allows
the sperm to get into the brood pouch and fertilize the egg is
unclear. It is quite possible that the jerks of the female’s
postabdomen create an inward current that helps bring the
sperm into the brood pouch.”
It is possible that the elongated first antennae of
male daphniids and moinids are used to feel the roughened carapace of a female (Goulden, 1968; Kokkinn
and Williams, 1987). The female’s carapace ornamentation might tell the male whether she is the correct
species and that she is producing haploid eggs. As with
all aspects of behavior in Daphnia, we know few details of the individual behaviors that result in mating.
Instead, cladoceran behavior has been studied mostly
at the population level, using plankton nets in lakes.
B. Individual Swimming Behavior
Video and computer analysis techniques allow the
analysis of three-dimensional swimming of individual
pelagic cladocerans, especially Daphnia to measure
swimming speeds, directions of movement (Dodson
and Ramcharan, 1991). The characteristic hopping
motion has not yet been satisfactorily characterized.
Average swimming speed varies from just enough to
maintain position (all pelagic cladocerans sink if quiescent), to sustained average speeds as high as about
25 mm / s (Dodson et al., 1997b). Sinking rate is
affected very little by temperature, less so than predicted by changes in water viscosity, suggesting Daphnia have behaviors to maintain constant sinking rate
(Gorski and Dodson, 1996). Different clones swim differently, with the fastest swimming characteristic of
clones from low-predation habitats. This suggests there
is a benefit to fast swimming (perhaps increases filtering rate) that is sacrificed in communities with high
predation rates (where encounter rates can be minimized by slow swimming). Fish predators pay attention
to Daphnia swimming behavior, preferentially taking
faster swimming individuals of real prey (O’Keefe
et al., 1998) or virtual (computer-simulated) prey
(Brewer and Coughlin, 1996).
Predaceous cladocerans swim slowly through the
water. The slow movement is probably a compromise
between swimming fast enough to run into prey and
slow enough to avoid attracting the attention of fish.
Browman et al. (1989) describe the swimming movement of Leptodora (Fig. 60):
“Leptodora swims randomly [at about 13 mm per sec] through
the water column with all five pairs of thoracic appendages
spread to form a ‘feeding basket’ and, seemingly by chance,
encounters prey. Shortly after prey make contact with any part
of Leptodora’s body, (usually ventral), the abdomen is rapidly
pulled forward, clamping itself under the feeding basket so that
the telson closes it at the posterior end. The duration of this
movement is always the same and we conclude that it is an
indiscriminate reflex. If the prey is encountered anywhere but a
short distance directly in front and slightly below the
Leptodora, it is not captured.”
C. Diel Vertical Migration
One of the earliest observations of cladoceran behavior at the population level is Cuvier’s, 1817, description of Daphnia vertical migration (translated by
Bayly, 1986):
“In the morning and evening, and even during the day when the
sky is overcast, the daphnids usually stay at the surface. But
during very hot weather, and when the sun beats fiercely down
on the pools of stagnant water where they live, they sink down
in the water, and stay at a depth of six or eight feet or more;
often not a single one can be seen at the surface.”
Cladoceran populations have been observed to migrate, on a daily cycle in both fresh and marine waters
(Hutchinson, 1967; Bayly, 1986; Dodson, 1990). Distances migrated by pelagic species vary from less than a
meter in a small rock pool to hundreds of meters in
oceans. In freshwaters, the usual distance is 5 – 20 m.
Because lake waters are vertically stratified, vertical migration takes the animals into very different habitats in
terms of temperature, light, food, turbulence, oxygen
and salts, and predators (Landon and Stasiak, 1983).
The greatest depth of the migration is often set by the
21. Cladocera and Other Branchiopoda
bottom of the lake, by the depth to which light penetrates, or by the limit of sufficient oxygen (about
1.0 – 0.5 m␥ⲐL; Weider and Lampert, 1985; Nebeker
et al., 1992). Littoral organisms also have a pronounced diel up-and-down migration. This behavior
allows 90% of the population below a sample area to
be trapped over night in inverted funnels (Whiteside
and Williams, 1975).
The daily pattern often seems to have the greatest
amplitude during the summer and fall months. The
larger developmental stages typically migrate the farthest. Populations usually migrate downwards at about
dawn and do not return until after dusk.
Physiological response to light intensity and duration is a component of vertical migration behavior
(e.g., Ringelberg, 1987). Directionality of light source
is important for Daphnia to be able to assume normal
swimming position; if the light comes equally from
every direction, the animal is unable to swim normally.
Cladocerans can probably orient in the water using polarized light (Ringelberg, 1987). There is also some evidence that cladoceran swimming speed and direction is
influenced differently by red and blue light.
Many ecologists have concluded that, while response to light is an important element in vertical migration, the most important factor is escape from
predators (Lampert, 1993; DeMeester et al., 1998;
Tollrian and Harvell, 1998). Cladocerans move downward during the day, into the darkness of deep water,
where they are relatively safe from visual predators
such as fish. However, deep water also tends to be cold.
In order to grow and reproduce as fast as possible, the
cladocerans return to warmer water near the surface at
night. This story has been elegantly demonstrated by a
series of experiments in the large plankton towers at
the Max Planck Institut für Limnologie in Plön,
Germany (Loose, 1993). When exposed to a combination of light and feeding predators, or even the water
from cultures of predators, Daphnia rapidly swim
downward. They accomplish the vertical migration not
by sinking or gentle swimming, but by rapid directional
swimming (Dodson et al., 1997a). Because the prey respond to predator water, the behavior is clearly in response to a chemical signal released by the predator.
D. Swarming Behavior
Cladocerans sometimes form swarms. Baird (1850)
reported:
“I have, however, frequently seen large patches of water in
different ponds assume a ruddy hue, like the red rust of iron, or
as if blood had been mixed with it, and ascertained the cause to
be an immense number of the D. pulex. The myriads necessary
to produce this effect is really astonishing, and it is extremely
865
interesting to watch their motions. On a sunshiny day, in a large
pond, a streak of red, a foot broad, and ten or twelve yards in
length, will suddenly appear in a particular spot, and this belt
may be seen rapidly changing its position, and in a very short
time wheel completely round the pond. Should the mass come
near enough the edge to allow the shadow of the observer to
fall upon them, or should a dark cloud suddenly obscure the
sum, the whole body immediately disappear, rising to the
surface again when they have reached beyond the shadow, or as
soon as the cloud has passed over.”
Swarms and movements similar to those described
by Baird (1850), if not as colorful, have been reported
for other planktonic and littoral cladocerans
(Weismann, 1880). Tessier (1983) described swarms of
Holopedium, and reviews reports of other cladocerans,
including Ceriodaphnia, Bosmina, Daphnia, Moina,
and Sida. Daphnia and Polyphemus sometimes form
small swarms of a few hundred animals. Although earlier studies with Ceriodaphnia, Daphnia, and Moina
suggested that females in swarms tended to produce
ephippial eggs, Crease and Hebert (1983) failed to find
any evidence for sexual pheromones produced by either
sex of swarming Daphnia. Butorina (1986) observed
Polyphemus swarms consisting of a few hundred animals in mushroom-shaped swarms about 50 cm deep,
6 – 30 cm in diameter, and with most of the animals in
the top 5 cm of the swarm at the surface of the water
(Butorina, 1986). The swarms were simple aggregations, with no difference in age structure from the surrounding nonswarming Polyphemus. The swarms are
dynamic, with animals arriving and leaving as long as
the sun shines. In addition to light, fish smell is also an
environmental signal that appears to induce swarming
behavior (Jensen et al., 1998). Compared to controls,
Daphnia in fish water swim more uniformly.
Ilyocryptus (Fig. 44) has a reputation for nesting
(Williams, 1978). They are often found in dense aggregations, which have been interpreted as the result of
young staying in the vicinity of their parent.
E. Escape Behavior
Cladocerans show various escape behavioral responses to predators (reviewed in Brewer et al., 1998).
At least some of the planktonic species have a fastswimming response which takes them away from
predators. This is effective against nonvisual predators
such as Leptodora (Browman et al., 1989), but may be
useless against fish. Diel vertical migration or small
body size are the most effective defenses against large
predators, such as fish, that use vision to locate prey.
When caught by a small predator, such as a copepod or
insect larva, cladocerans struggle to break free.
Bosmina, once it escapes a nonvisual predator, closes
up its carapace and sinks without making detectable
866
S. I. Dodson and D. G. Frey
hydrodynamic pressure waves (noise) (Kerfoot et al.,
1980). Also, Bosmina, like chydorine chydorids, tucks
the second antennae under the carapace for further
protection when captured.
Brewer et al. (1998) measured the three-dimensional escape response of Daphnia. Daphnia of a given
size always escaped at the same speed, regardless of
origin of the clone, but individuals from lakes with fish
tended to start escaping much sooner than those from
ponds lacking fish.
V. WHAT CLADOCERANS EAT
AND WHAT EATS CLADOCERANS
Planktonic cladocerans, such as Daphnia, are of
great importance to the ecology of lakes, affecting the
amount of algae, via grazing and nutrient regeneration,
and providing food for young-of-the-year fish (Carpenter, 1987; Kitchell, 1992; Sommer, 1989; Lampert and
Sommer, 1997).
A. Cladoceran Feeding
Cladocerans are discriminating in their choice of
food (Porter, 1977). There are two major aspects of
foraging relationships: what cladocerans eat, and how
they get it. Techniques recently developed allow quantitative measurements of the various aspects of the feeding process: food collection, ingestion, assimilation,
and defecation (Peters, 1984).
The so-called “herbivorous” cladocerans eat a
variety of small particles, from bacteria less than a
micrometer long (Porter et al., 1983), through algae,
up to ciliates, small rotifers, and copepod nauplii as
large as 100 m (Porter, 1973; Burns and Gilbert,
1986; Dodson, 1975; Pace et al., 1994). The most important component of their diet is made up of small algae in the range of 1 to 25 m. Algae larger than about
50 m or algae with spines or in colonies are usually
rejected. Also, cyanobacteria in the preferred size range
are often toxic and are not eaten if abundant (Porter
and Orcutt, 1980). The size of food particles is determined by more factors than the body size of the herbivore. For example, Bosmina, a small filter feeder, specializes on larger particles than do larger Daphnia
(DeMott and Kerfoot, 1982).
Chydorids typically feed by crawling along surfaces or through mud and scraping up or filtering food
(Fryer, 1968; Smirnov, 1971). Many cladocerans use
the carapace to help filter feed or as a pair of cheeks
which confine the feeding current. However, some
chydorids also employ the carapace as a suction cup.
Negative pressure can be created inside the carapace if
water is pumped out, as long as there is a good seal between the carapace and substrate. Second antennae,
which are used for swimming by pelagic cladocerans,
are used for crawling and jumping in some chydorids.
Thus, chydorids have a number of specialized feeding
and related locomotion adaptions. For example, Eurycercus, the largest chydorid, climbs on upper surfaces of
plants and is able to filter feed. Alonopsis elongata balances upright on its carapace margins on solid surfaces
and scrabbles along using its second antennae, scraping
up living and dead organic material with its thoracic
legs. Peracantha truncata keeps its second antennae inside the carapace when it crawls and can crawl upsidedown if the surface is rough enough to provide a hold
for the claws on the first leg, aided by weak negative
pressure. Alonella exigua can hang upside-down on
aquatic vegetation solely by means of negative pressure. Graptoleberis testudinaria has the most effective
pressure chamber and exhibits snail-like behavior. Leydigia leydigi has neither pressure chamber nor scraping
spines on the thoracic legs. It pushes itself into soft
ooze using its first antennae and large spiny postabdomen. Food is sieved from the mud using setae along
the margin of the carapace and on the shortened thoracic legs. In Leydigia the respiratory current (which
passes over the outside of the first two thoracic legs) is
separated from the feeding current. While most chydorids are herbivorous, the scraping method of gathering food lends itself to carnivory, and Anchistropus is a
predator on hydra, while Pseudochydorus globosus is a
scavenger of dead animals.
Leptodora, Polyphemus, and Bythotrephes are
predaceous cladocerans, eating protozoa, rotifers,
planktonic stages of small chironomid larvae, and
small crustaceans. Leptodora takes prey between about
0.5 and 1.5 mm and seems to prefer other cladocerans
(Karabin, 1974; Browman et al., 1989). The smaller
polyphemids take smaller prey and prefer small cladocerans, rotifers and protozoans (Monakov, 1972;
Lehman, 1987). Algae is a minor component of the diet
of Polyphemus and Bythotrephes. Presumably, the
metanauplii of Leptodora eat algae. Adult Leptodora
and Onychopoda have a reputation of being vampirelike “fluid feeders” (Cummins, 1969; Karabin 1974).
Leptodora (and other predaceous cladocerans) need actual contact with prey before they attempt to capture it
with the outspread thoracic legs (Browman et al.,
1989). Once the prey is grasped, the Leptodora bites it
with the mandibles, sucking up body fluids and perhaps small bits of tissue. Small prey, such as rotifers
and protozoans are probably chewed and then swallowed. Edmondson and Litt (1987) report large numbers of whole rotifers (Conochilus) and even whole cyclopod copepods in the gut of Leptodora.
21. Cladocera and Other Branchiopoda
B. Predators on Cladocerans
Cladocerans are eaten by any freshwater predator
that can either swallow them or chew pieces out of
their small bodies. A few examples of predators include: common pond insects (Cooper, 1983), especially
odonates (Hirvonen and Ranta, 1996) and larvae of
the phantom midge Chaoborus (Swift and Federenko,
1975), flatworms (Schwartz et al., 1983), hydra (Havel,
1985), salamanders (Dodson and Dodson, 1971), fish
of all kinds but especially small pelagic forms (O’Brien,
1979) except for the smallest immature fish (Graham
and Sprules, 1992), birds (Dodson and Egger, 1980),
predaceous copepods (Dodson, 1974; 1984; Havel,
1985), other predaceous cladocerans (Havel, 1985),
and even predaceous plants (Havel, 1985).
Examination of the gut contents of minnows, sticklebacks, and small fingerlings of other fishes shows a
heavy uptake of littoral cladocerans. The young of percid and centrarchid fishes moving through the macrophyte beds in the littoral zone find an abundance of
these animals to consume. Small catostomid fishes in
streams near Bloomington, Indiana seem to feed almost
exclusively on chydorids. Whiteside and his students
believe that one of the possible causes of the midsummer population crash of chydorids in lakes of northern
Minnesota is predation by early fingerling perch. The
small size of the littoral cladocerans is suited to the
feeding abilities of small fishes.
VI. POPULATION REGULATION
Cladoceran populations, especially of the planktonic species, are characterized by hundredfold annual
variations in population size. The peak population
generally occurs during an algae bloom, which occurs
when lakes are mixed in the spring in northern areas or
when the rainy season begins in southern areas. Cladoceran populations are typically at low points during
cold weather or when edible algae are scarce. Littoral
species show similar population dynamics (Whiteside,
1978).
What causes the large variations in Daphnia abundance? Until the work of Hall (1964), it was assumed
that the only significant factor was the availability of
edible algae. However, Hall showed that while the increase in Daphnia numbers was due to an algal bloom,
the decrease was due to predation rather than starvation. Recent studies suggest that the usual cause of the
decline in abundance is often a combination of predation and food limitation (Lampert, 1986b). Daphnia
possess enough energy stores to carry them through
short-term food shortages, such as the “clear-water
867
phase” that sometimes follows peak Daphnia abundances. However, lowered fecundity during the summer,
after the clear-water phase, indicates food limitation.
Mortality due to predation is generally sizespecific, depending on the size of the prey and the size
and hunting behavior of the predators (Zaret, 1980).
Those cladocerans that can coexist with predators tend
to have specialized life histories, morphologies, and behaviors that provide defenses against predation. Even
so, mortality due to predation can be an important factor in determining the abundance of zooplankton.
Hall (1964) made a great contribution to aquatic
ecology by showing that predation, as well as food limitation, limits zooplankton populations. He did this by
separating the population growth rate coefficient (r)
into a component of birth or recruitment (b) and a
component of mortality (d). He used a life table technique which requires that individual animals be cultured in the laboratory to determine rates of mortality
in the absence of predation and fecundity at various
food levels. This important process of separating mortality and recruitment can be achieved much more easily using Edmondson’s (1960) egg-ratio technique. This
technique requires that one know the number of eggs
per female and the developmental time of an egg in order to estimate b. Then, r can be estimated from
changes in population size, and d is estimated by the
difference between b and r. This technique provides
good estimates of b when compared to life table estimates (Lynch, 1982), even when the assumptions of the
egg-ratio technique (concerning age structure) are not
met. The original Edmondson egg-ratio technique has
been modified in various ways (Rigler and Downing,
1984). A comparison of these modifications with predictions based on life table studies (Gabriel, 1987) suggests that different modifications are most accurate depending on the population age structure and the
frequency with which the population is sampled.
In general, a declining population with a high
value of b indicates high predation, and a stable or declining population with a low value of d indicates food
limitation. Studies on the effects of both food limitation and predation suggest that both regulate populations. The best competitors are generally the most
susceptible to predators. In a lake dominated by Holopedium and Daphnia, the Holopedium were mainly
limited by the food supply, while Daphnia showed significant mortality from fish during the summer (Tessier,
1986). A combination field and laboratory study
showed that Holopedium required more food than
Daphnia. Thus, when fish predation was intense and
food was abundant, Holopedium increased; but whenever food was scarce, Holopedium declined relative to
Daphnia; and when predation was weak and food
868
S. I. Dodson and D. G. Frey
abundant, Daphnia increased faster than Holopedium.
Vanni (1986), using large plankton enclosures in a eutrophic lake, found that large Daphnia could reduce
food levels sufficiently to reduce populations of
Bosmina and several copepods. The Daphnia in the
lake would have been kept in check by fish predation.
Cooper and Smith (1982) found that when fish are absent, large invertebrate predators such as Buenoa could
function in a size-selective manner like fish and remove
the larger Daphnia pulex, which was able to live at
lower food concentrations than the smaller Daphnia
laevis. Kerfoot and Sih (1987), Carpenter (1987), Sommer (1989), Kitchell (1992), Lampert and Sommer
(1997) provide further examples of direct and indirect
effects of predators on cladoceran abundance.
VII. FUNCTIONAL ROLE IN THE ECOSYSTEM
Hrbácek (1958) first pointed out the importance of
certain fish in excluding cladocerans from communities. These ideas were extended to the effects of selective grazing of cladocerans on algae to produce a
model of a dynamic food chain, in which the number
and abundance of species in a community is the result
of the interaction of competition, selective predation,
and nutrients (Fig. 6). For many aquatic predators,
cladocerans are a major diet item.
While phosphate concentration in a lake is a major
determinant of the kind and abundance of algae, and
hence of “water quality,” the phosphate concentration
is not a particularly good predictor of water quality
(Schindler, 1977). For any given level of phosphate,
there is great variation in water quality. Several recent
studies suggest that food chain dynamics also determine water quality (Shapiro, 1982; Carpenter, 1985;
McQueen, 1986; Carpenter, 1988; Kitchell, 1992).
Large species of Daphnia appear to be the most efficient in cleaning up lakes made turbid by small algae.
Filter-feeding cladocerans can remove significant fractions of the algae from a lake each day (Porter, 1977).
Each animal, depending on its size, can clear 1 mL
per animal per hour (Porter et al., 1982). For example,
large Daphnia pulicaria was scarce in Lake Washington
until 1976, when it became common (Edmondson and
Litt, 1982; Edmondson, 1991). At the same time the
mean summer transparency of the lake doubled, due to
grazing by Daphnia on diatoms and other algae. The
large size and high feeding rate of the large Daphnia
make them especially efficient at cleaning up lakes.
The hypothesis of food-chain dynamics is a central
hypothesis in ecology, because of the perspective it provides and the potential it has for explaining the diversity
of natural systems (Fretwell, 1987). The power of this
hypothesis results from its inclusion of both direct effects, such as predation of fish on zooplankton, and indirect effects, such as the effects of fish predation on the
abundance of algae eaten by zooplankton. In aquatic
communities, this hypothesis predicts that the abundance of algae (especially nuisance algae, such as the
scum-forming blue green algae) can be kept at low levels by maintaining dense zooplankton populations,
which depend on the scarcity of plankton-eating fish,
which in turn are controlled by an abundance of large
fish-eating fish. Thus, the theory of food-chain dynamics predicts that management of game fish, such as bass
or pike in a lake, will have effects on the abundance of
algae and, consequently, various aspects of water quality depending on algae, such as clarity and odor. A key
link in this chain are the large algae-eating zooplankton,
especially Daphnia, which are common and often abundant, the favorite food of plankton-eating fish, and able
to eat rapidly large quantities of algae (Carpenter,
1985). Smaller common cladocerans, such as Bosmina
and Ceriodaphnia, while less susceptible to fish predation, are also less efficient at cleaning up algae.
Meiobenthic (littoral) cladocerans constitute one of
the prime converters of organic detritus with its associated microorganisms into particles big enough to be
handled by small fishes and predaceous invertebrates.
Wetzel (1983) demonstrated that often the littoral community is more important than the open-water community in the annual production of a lake.
VIII. PALEOLIMNOLOGY AND MOLECULAR
PHYLOGENY
A. Paleolimnology
In paleolimnology, we seek to interpret how a lake
functioned in past time. The sediments accumulate
chronologically; and hence from changing representation of kinds of sediments and their sorting coefficients,
organic and inorganic chemicals, and morphological
remains of plants and animals, we attempt to interpret
past conditions in the water bodies and their watersheds. Sometimes the record is exceptional. Where
varves (annual layers) are present, changes in response
from year to year can be determined.
Lake sediments tell fascinating stories about the
lake and the watershed. Lake sediments have been used
to understand the past climate (from pollen from terrestrial plants in the watershed), lake productivity
(from the kinds and amounts of microfossils of lake organisms), and the effects of species invasions into the
lake. Lake sediments can also supply important archaeological insights. For example, a study of a small lake
21. Cladocera and Other Branchiopoda
in central Italy (Lago di Monte Rosi; Hutchinson et al.,
1970) found a close correspondence between the 14C
date for a thin layer of charcoal and actual records of
Roman road-building activity in the watershed.
Remains of animals that are preserved in lake sediments are mostly either silicious (sponges), calcareous
(ostracods), or chitinous (cladocerans, midges). All
groups of freshwater animals leave at least some remains in sediments (Frey, 1964), but certainly the
cladocera are usually best represented, with quantities
of remains generally between a few thousand and several hundred-thousand per cubic centimeter of fresh
sediment. Once the species pool of a region is known,
then virtually all the fragments can be identified positively as to producing species. Remains of the chydorids and bosminids are preserved best and nearly
quantitatively, but all cladoceran taxa leave some remains. Those of the meiobenthic forms become integrated over time (at least a year) and habitat before being incorporated into deepwater sediments. The
quantity of these remains varies from place to place,
being greater toward shore than in the profundal zone,
but the percentage composition by species is the same
all over the bottom (Mueller, 1964). The best way to
determine what species of cladocera occurs in a water
body and in what relative abundance over time is from
the animal remains in the sediments.
Close-interval stratigraphies are constructed of
these remains in the same way as for pollen or diatoms,
yielding changes in relative abundance over time of the
various taxa (Frey, 1986b). Interpretation has been difficult because of an insufficient knowledge of the ecology of these forms. A few species, such as members of
the Chydorus sphaericus complex, are considered to respond positively to eutrophication, sometimes comprising more than 90% of all remains. Others seem associated with bog conditions or changes resulting from
acidification. Daphnia and Bosmina fossil remains
record past predation regimes (Kerfoot, 1981; Kitchell
and Carpenter, 1987).
What is really necessary is to analyze the community differences between time and place. Cotten (1985)
studied cladoceran communities in 46 lakes in eastern
Finland, using their remains in surficial sediments. She
obtained about 70 different taxa. These and their relative abundances were then ordinated using measured
chemical and physical characters of the water bodies
and their watersheds. She recognized five kinds or
groups of lakes that were almost completely concurrent
with the five kinds revealed previously by diatom
analysis. Thus, the cladocera are just as good responders as diatoms to conditions in a water body and to
changes in these conditions over time naturally or in response to human activities.
869
A study of the cladocera present in a lake in Denmark during the last interglacial period (roughly
100,000 years ago) revealed 25 species of chydorids,
all of which occur in Denmark today (Frey, 1962). No
real differences in morphology were apparent; and
moreover, the relative order of abundance of these 25
species in the interglacial was the same as in Denmark
today, indicating that the ecology of these forms had
not changed either.
Furthermore, evidence and suggestions from the
meager remains of cladocera known from the present
to the Tertiary, Cretaceous, and possibly even the Permian indicate that major evolution in the cladocera had
occurred by the late Paleozoic or early Mesozoic, and
that the species present today have been stabile for millions of years. Thus, knowledge of the present ecology
of species will probably be usable in interpreting relationships in lakes even in the distant past.
The presence of a vast record of cladocera in lake
sediments is valuable for more than just paleolimnology.
Sometimes the presence of a particular species of cladoceran at a place is suspected to be the result of establishment by recent transport there, such as Bythotrephes in
the Great Lakes (Sprules et al., 1990). This hypothesis
can be checked by sedimentary analysis if desired.
The development of the cladoceran community in
a lake since late-glacial time, or even earlier in
nonglacial lakes, can be followed easily (Goulden,
1966). The species diversity increases over time as the
number of taxa in the community increases as conditions become stabilized, tending toward the limit for
maximum diversity at any given number of taxa
(Goulden, 1969). The distribution of species by their
relative abundance in a stable community closely follows the MacArthur broken-stick model. Any major
deviation in fit to the expected is considered to have
arisen from changes in climate, agriculture, volcanism,
or other controlling agents (Frey, 1974, 1976). In lakes
with varved sediments one can follow changes in relative abundance of the total cladoceran community on a
year-to-year basis and then relate these changes to potential causative agents (Boucherle and Zulig, 1983).
Access to the literature of paleolimnology can be
obtained from the proceedings of the four international
symposia on the subject (Frey, 1969; Klekowski, 1978;
Löffler, 1987; Merilainen, 1983; North American Lake
Management, 1995).
B. Cladoceran Fossil Record
Fossils of branchiopods and their relatives are
known from the Cambrian Period, about 530 million
years ago. Kerfoot and Lynch (1987) summarized data
on the earliest fossils of Branchiopods. In the Cambrian
870
S. I. Dodson and D. G. Frey
FIGURE 5
(A) Phylogenetic tree, showing the divergence of cladoceran genera during the Mesozoic, and
differentiation of the subgroups of Daphnia during the Cretaceous (Lehman et al., 1995); (B) phylogenetic tree
showing the species in the three major species groups of the genus Daphnia (Colbourne et al., 1997).
21. Cladocera and Other Branchiopoda
Period, animals first developed mineralized coverings
about 540 million years before the present (mybp).
There are branchiopodlike fossils from the Burgess
Shale, such as Protocaris and Branchiocaris, but these
animals are not considered branchiopods (Gould,
1989). A fossil of an animal that is probably an ancestor of the branchiopods (and distinct from other crustaceans) occurs in Upper Cambrian sediments of Sweden (Walossek, 1995).
By the lower Devonian (ca. 410 mybp, when fish
also make their appearance), the larger Branchiopoda
are well established as three recognizable modern
groups: fairy, tadpole, and clam shrimp. The fairy
shrimp are represented by Lepidocaris, an anostracanlike animal, but with a long segmented second antenna
and more mouth parts than modern fairy shrimp. These
animals are from sediments that indicate marine and estuarine, and perhaps freshwater sediments. Tadpole
shrimp fossils are found in marine sediments, and a
clam shrimp fossil also appears in marine sediments
(this animal is possibly an ancestor of both modern
clam shrimps and cladocera). By the Carboniferous (ca.
286 – 360 mybp), tadpole and clam shrimp species are
diverse and occur in many habitats (marine, freshwater,
streams). Smirnov (1970) described some possible
Daphnia-like and chydoridlike fossil from the late
Permian (ca. 240 mybp). The Daphnia-like animals are
5 – 8.5 mm long. Smirnov also has good specimens of
Daphnia-like fossils from the Jurassic (ca. 225 mybp).
By the early Triassic, most tadpole and clam shrimp
groups are extinct, and cladocera appear, perhaps as
neotenic conchostracans (as suggested by Brooks). The
modern continental distribution of cladoceran orders
suggests the orders were distinct when the large continent Pangea broke up into Laurasia and Gondwanaland, during the Cretaceous (approximately 100 mybp).
Starting with the Oligocene (ca. 38 mybp), there are
many good fossils of cladocerans, especially of ephippia
from lake deposits such as the freshwater Florrisant
beds in Colorado. Similarly, the modern distribution of
cladoceran species in North America and Europe suggests that the major about of speciation associated with
glaciation was associated with speciation which
produced modern species (Brooks, 1957). Continental
glaciers have been producing new lakes since about
1.8 mybp. For example, several species, such as Daphnia galeata, retrocurva, catawba, longiremis, are restricted to the glaciated portion of North America (or
Europe: D. cuculata).
C. Molecular Phylogeny
Just as lake sediments contain a record of the
history of the lake and its watershed, the nucleic acid
871
(nuclear and mitochondrial) of cladocerans contains a
record of the evolutionary history of living individuals.
The molecular record is particularly valuable, because
the fossil record of most zooplankton groups is poor.
Information about phylogeny exists in the nucleic
acids of branchiopods. For example, analyses of the ribosomal RNA gene (encoded on mitochondrial DNA)
suggest that Daphnia originated (differentiated from
similar genera) in the Mesozoic (Lehman et al., 1995;
Colbourne et al. 1997; see Fig. 5A). Since then, the
genus has evolved into at least three species complexes
(Fig. 5B). A recent re-analysis of an Australian daphniid (Daphnia jollyi), suggests it may represent a fourth
lineage. It is odd that there are only four complexes of
Daphnia species, and that they are all rather similar
morphologically, given 200 – 300 million years of evolution.
IX. TOXICOLOGY
Cladocerans, especially Daphnia, Ceriodaphnia,
and Moina have long been used in toxicity tests. Cladocerans are relatively small, have short life spans, reproduce well in the laboratory, and are ecologically important — all desirable characteristics for a toxicity test.
For a bioassay (a test using organisms), the animals are
grown under optimal conditions in the laboratory (or
less often, in enclosures in nature). Some animals are
treated with known concentrations of chemicals, and
their responses are compared to those of untreated animals. Survival and fecundity (number of offspring) are
two standard bioassay endpoints (US EPA, 1994). Animals are exposed to test chemicals for a few hours or
days as a test for acute toxicity, and for a generation or
more as a test for long-term chronic toxicity.
Morphology, behavior, sex ratio, physiology, and
developmental time are additional bioassay end points.
These end points tend to be more sensitive than survivorship or fecundity, and they are of ecological importance. For example, the insecticide carbaryl induces
morphological and swimming behavior changes in
Daphnia (Hanazato and Dodson, 1993; Dodson et al.,
1995). The surfactant (nonionic detergent) and plasticizer nonylphenol interferes with normal development
at levels found in streams and lakes, so that neonates
are produced and they survive in the laboratory, but
they are unable to swim (Shurin and Dodson, 1997).
Predator prey behavior is affected at low-dose and
sublethal levels of many environmental pollutants. Carbaryl causes Daphnia to swim erratically and faster
than normal (Dodson et al., 1995), which in nature,
would increase the predator – prey encounter rate, leading to an increased Daphnia mortality.
872
S. I. Dodson and D. G. Frey
FIGURE 6
Food-chain models showing: (A) four trophic levels; (B) three trophic levels; and (C) three
consumer trophic levels and one scavenger (decomposer) level.
21. Cladocera and Other Branchiopoda
Many aspects of animal development are controlled by hormones. There is concern that industrial
and agricultural chemicals may be acting like or interfering with normal hormone systems in both wildlife
and humans (Colborn et al., 1996). The cladoceran sex
ratio is an endpoint that can be used to test for these
chemicals (called “endocrine disruptors). Chemicals
that change sex ratio in cladocerans will have ecological consequences, and they may also act as endocrine
disruptors for other animals, including humans. It is
hypothesized that some chemicals that disrupt hormone systems in vertebrates can also interfere with the
system regulating molting in arthropods (Zou and
Fingerman, 1997).
X. CURRENT AND FUTURE RESEARCH
How many species are there of cladocerans? Do
cladoceran species need conservation? How do you
identify the different species? How closely related are
the different morphologically similar species? Are
cladocerans (or other branchiopods) even really crustaceans? There is a great deal of taxonomy and phylogeny that remains to be done. This is not just a matter of finding all the species and naming them, but
more fundamentally, of deciding what sort of species
concept makes sense with cladocerans. These animals
are usually at least facultatively asexual; and in many
groups, what seem to be good morphological species
are made up of a combination of occasionally sexual
and totally asexual populations. Descriptions of species
before about 1950 are mostly very inadequate and
hence cannot be used to compare populations on two
different continents claimed to be the same taxon. To
determine that any North American taxon is the same
as or different than the taxon of the same name described from elsewhere is difficult. One first needs to
compare both taxa closely, using large populations
containing all instars and reproductive stages. Thus far
about 16 pairs of taxa with the same name, from the
New and Old World, have been compared (partial
summary in Frey, 1986a), with the finding that almost
all of them are full species. Sometimes, as in the
Chydorus sphaericus complex, Eurycercus, Ephemeroporus, the Chydorus reticulatus group, the Chydorus
faviformis group, and Pleuroxus laevis, the same specific name was being applied to two or more related
but different species in the same geographical region.
Consequently, for critical studies in ecology, one must
be careful not to transfer the ecological requirements
and indications of a species across an ocean or some
other distributional barrier, on the assumption that all
taxa presently bearing the same name are the same
873
species. This question will be sorted out with a mixture
of traditional morphological and new molecular and
genetic techniques (e.g., DeMelo and Hebert, 1994;
Colbourne et al., 1997), and some new ideas!
The study of food-chain dynamics, using sets of
whole lakes shows promise for interesting research into
population and community ecology. Society has a great
desire for clean lakes and lots of fish production. Cladocerans play crucial roles in both water quality and fish
management, but we need to understand cladoceran biology better before we can meet society’s desires. Food
chain dynamics, or “biomanipulation” shows promise
of being an inexpensive and safe method for improving
both water quality and fish production.
Zooplankton live in a sea of chemical signals (Larsson and Dodson, 1993). The chemicals are produced by
predators, food, competitors, and environmental sources.
The signals induce changes in development, morphology,
reproduction, and behavior. However, none of the chemical signals used between species have been characterized.
When their chemical structure is known, it will be possible to do interesting experiments involving induction,
and to understand how these signals affect development
and behavior, which, in turn, affect the aquatic community and trophic dynamics.
Environmental toxicology has long depended on
tests of cladoceran survival and fecundity. Future research will continue to use more ecologically relevant
endpoints, such as behavior, developmental time, and
sex ratio (Dodson et al., 1999).
XI. COLLECTING AND REARING
A. Sampling Techniques
Cladocerans can be collected using a fine mesh net
pulled through the water or a sieve through water is
poured. A large number of specialized nets have been
developed for specific applications (Bernardi, 1984).
The mesh should be around No. 10, or have a pore size
of about 90 – 150 m. Nets are most useful for towing
in open water. Weedy habitats can be sampled by a
plankton net attached to a handle, and with a widemesh (1 cm or so) cover over the opening of the net.
Weedy habitats or mud can be sampled by picking up
water in a bucket and pouring it through a series of
nets or sieves (Downing, 1984). In any case, keep mud
out of the sample if possible. Large pieces of water
weeds and detritus can be filtered out of a sample with
screen having a mesh size of about 1 cm. Filters can be
made by cutting the bottom off a plastic bottle. The
center of the cap is cut out, and a piece of mesh glued
over the hole. When the cap is screwed back onto the
874
S. I. Dodson and D. G. Frey
bottle, water can be poured into the inverted bottle,
and cladocerans will be caught by the mesh. The captured cladocerans can then be washed into a jar of water previously filtered through the mesh.
Estimations of population dynamics or productivity require quantitative samples, which require special
gear (Bernardi, 1984; Downing, 1984) and a careful
experimental design and statistical analysis of samples
(McCauley, 1984; Prepas, 1984).
If the sample organisms are to be killed, there are
at least two ways to do it (the authors disagree on the
best technique). Dodson prefers to wash the animals
off the screen of a small sieve using a wash bottle filled
with 95% ethanol, or to drop the sample into a large
volume of 95% ethanol. This will kill the animals
rapidly enough so there will be little distortion and will
preserve the sample indefinitely. It will be necessary to
keep the samples tightly capped and to add 70% alcohol from time to time. Although messy, a little glycerine will ensure that the sample never dries up. Frey preferred to kill and preserve animals by adding enough
commercial formalin containing 40 g sucrose per liter
to make a 5% formalin solution. He felt formalin was
better for long-term museum preservation of cladocerans, because of lesser distortion of the specimens over
time. For safety reasons, formalin samples are best examined in plain water, following filtration. Specimens
can then be replaced in a formalin solution for storage.
B. Culture Methods
Animals will live in filtered pond water for a few
days at least, especially if there are fewer than 50 animals / L. Animals can be transferred to new water using
pipettes made of glass tubing and rubber bulbs, or for
large quantities, turkey basters are useful. For longterm studies of cladocerans, it is possible to maintain
cultures for years. Maintenance of cladoceran cultures
can be divided into two processes: maintenance of food
for the cladocerans and care of the cultures.
1. Cladoceran Food
The simplest to maintain and most reliable bulk
food culture for cladocerans appears to be a mixture of
green algae and bacteria maintained in a large aquarium. Keep the algae in a glass aquarium as large as possible; a 40-gal aquarium is sufficient. Place the aquarium in a south-facing window. Choose a place that can
be kept at least 18 – 25°C; a greenhouse is preferable.
Fill the aquarium with filtered lake or pond water, using
a fine mesh (ca. 90 m mesh) net, or use tap water aged
for 1 day with a little additional filtered lake water as a
seed source for the phytoplankton. Add a dozen or so
small fish (bait fish, guppies, or gold fish) to the tank
and feed the fish regularly. Aerate the tank. Keep the
glass walls scraped clean and siphon off bottom detritus
now and then. Regardless of what you start with, the
algae species will probably eventually stabilize, and you
will have a mixture of small greens (like Chlorella, Selenastrum, and Scenedesmus), along with a few other less
common species such as Ankistrodesmus. Keep the algae in a healthy state by removing at least a quarter of
the culture every week or so. Use a large-diameter
siphon to remove the green algae.
Well-equipped laboratories can grow single-species
algal cultures purchased from biological supply houses.
Grow the algae either in a medium supplied by the supply house, or mix artificial lake water, such as “Combo”
(Kilham et al., 1998). The advantage of Combo is that it
has the same salinity as average lake water, and the algae
can be added directly to branchiopod cultures. Grow the
algae in 2-L clear plastic bottles (soda bottles). For fast
growth, aerate the cultures and use bright light.
Culture Care: Branchiopods can be captured using
plankton nets, dip nets, or by pouring water through filters of about 100-m mesh. Wash the collected animals
into some filtered water from the lake or pond. Bring
the animals into the laboratory and let them equilibrate
to the laboratory temperature. Many cladocerans tend
to be captured by the surface film. This happens more
often for the smaller lake animals, especially if they are
initially in cold water with saturated oxygen, because
bubbles form on the carapace as the water warms. In
extreme cases of mortality, keep the surviving animals
in a narrow-mouth bottle filled to the top and plugged
with cotton pushed below the water surface. In the beginning, put the cultures on a dark substrate and keep
them in a room with dim light. After a couple of days, it
will be obvious which cultures are going to be vigorous.
Be sure to label the cultures (with tape on the jar) as to
the source and date of capture.
Feed the cultures algae suspension. If you cannot
see large objects through the culture jar, the algae is
probably too concentrated; dilute it. The mixture
should be green but not opaque. As the water in each
jar clears, add algal suspension. Do not let the water
become clear. Even an overnight bout of clear water
will cause some clones to starve. It is also possible to
overfeed clones, but starvation seems a much more
common occurrence. Continue adding algae suspension
until the jar is full.
After the jar is full, there are two ways to feed the
culture. First, if there are too many animals in the jar,
pour out half or so of the culture and refill with algal
suspension. Beware of cultures in which the animals
concentrate in the top centimeter or so of the water; it
is possible to pour out the entire culture! Second, if
there are too few cladocerans in the jar, siphon off
21. Cladocera and Other Branchiopoda
about half of the water and add algal suspension. Be
sure to use fine enough mesh (about 90 m) on the end
of the siphon in the jar, so that the animals stay in the
jar. Also, be sure to wash the siphon and dry it or dip it
into hot water before using it on the next culture. The
safest population size for a 1-L jar seems to be about
20 – 50 animals. Having some detritus on the bottom of
the jar will not harm planktonic species and is beneficial to meiobenthic species. When there is a thick layer,
decant off the culture and discard the detritus layer.
Clones, once they are established, can be kept in a
cold room or refrigerator at about 4°C. At this temperature, the animals need to be fed only about once every
two weeks. Beware — some species die when moved
from 20 to 4°C, so do not put all your cultures in the
cold at once.
XII. TAXONOMIC KEYS FOR
CLADOCERAN GENERA
A. Available Keys and Specimen Preparation
Crustaceans are cladocerans if they have four to
six pairs of (thoracic) legs, lack any paired eyes, swim
with their second pair of antennae, and have at least
the head not covered by a carapace.
The identification keys are based on a large number of sources, which are referenced in Table 1 and in
the individual keys. Dodson is responsible for most of
the keys. The key to the Chydoridae family was written
by Frey.
These keys are designed to allow identification of
adult females only. Dissection is seldom necessary, but it
is a good idea to have several animals available for identification, because these small, delicate animals are easily
crushed out of recognizable shape. Animals shorter than
about 1 mm should be mounted on a slide for highmagnification viewing; the larger species can often be
identified without being mounted, using a dissecting microscope at about 50 diameters magnification or less.
While live animals trapped in a thin film of water
can usually be identified, it may be necessary to kill
them with alcohol and mount them on slides. The best
mounting medium is one which is more or less permanent and which is soluble in water and alcohol solutions so specimens can be added directly to the medium
without tedious dehydration. If you have access to
chloral hydrate (currently a “controlled substance”), a
good medium is Hoyer’s:
875
Dissolve 30 g of ground Gum Arabic in 150 mL of
hot distilled water. Add 200 g of chloral hydrate
and 20 g of glycerine.
Divide the Hoyer’s into two batches. Let one sit in a
warm place until it is as thick as honey, and keep the
second batch in a closed jar so it remains thin. You can
always add more water to thin either batch. Keep at
least some of the thick and thin Hoyer’s in screw-top
eye dropper bottles for easy use, and be careful to keep
the Hoyer’s off the threads of the bottles.
To make a slide, use the eye dropper to put a small
streak of dilute Hoyer’s (about a quarter of a drop) toward one end of the slide. Arrange the specimens in this
streak. It is wise to place four or so of what you think
are the same kind of animal on a slide to allow for comparison and viewing of difficult characters. Delicate
specimens can be protected from compression by
putting a few pieces of broken cover slips around the
specimens. Let the Hoyer’s streak dry on a slide warmer
or in a warm place (below 100°C). When the streak is
dry, add a drop or two of the thick Hoyer’s and gently
lower a cover glass over the Hoyer’s. Put the slide back
into a warm place to dry again. Label the slide as to the
date and location of the collection! It is important to
use thickened Hoyer’s in the last step; otherwise, the
Hoyer’s will shrink as it dries, and produce large bubbles in the final preparation. On the other hand, if you
use thickened Hoyer’s, small bubbles produced when
you lower the coverslip will disappear as the medium
dries. These slides are semipermanent. If the climate is
humid, the Hoyer’s will thin and cover glasses will slip
off the slide. If the climate is arid, the Hoyer’s will desiccate enough so that the cover glasses pop off the slide.
These problems may be avoided by storing the slides
horizontally in a climate that has moderate humidity, or
you can ring the slide with nail polish.
An alternate mounting method, especially useful
for bosminids, chydorids, and macrothricids, is to use
polyvinyl lactophenol stained with lignin pink. Temporary slides can be made using glycerol or glycerine jelly.
Best of all for museum specimens is possibly Canada
Balsam, although this requires the specimens be dehydrated through an alcohol series.
B. Taxonomic Key to Families
of Freshwater Cladocerans
This key to families is followed by keys to genera
in each family or group of families.
1a.
Both branches (or branch) of second antenna ending in three setae (Fig. 1A) .....................................................................................2
1b.
At least one of the second antenna branches ending in four or more setae (Fig. 8) .............................................................................8
FIGURE 7 Holopedium gibberum Zaddach, 1855. Why Not Bog, Vilas Co., Wisconsin. 22 September 1973.
(KE) FIGURE 8 Sida crystallina (O.F. Mueller, 1776). Lake Mendota, Dane Co., Wisconsin. 11 September
1987. (KE) FIGURE 9 (A) Diaphanosoma birgei (Kořínek, 1981). Lake Wingra, Dane Co., Wisconsin. 5
October 1978. (KE) (B) Penilia avirostris Dana, 1849. Anteriolateral view. Atlantic Ocean near Florida
(29°55.8’ N and 81°07.4°W). 27 October 1987. (KE); (C) P. avirostris. Posteriolateral view. (KE) FIGURE 10
Pseudosida bidentata Herrick. 1884. Bull Frog Pond, S.R.P. area, South Carolina. 27 May 1985. (Coll. DGF
#7483). (KE) FIGURE 11 Latona setifera (O.F. Muelller, 1776). Lake Mendota, Dane Co., Wisconsin. 30
June 1987. (KE) FIGURE 12 (A) Latonopsis occidentalis Birge, 1891. Dick’s Pond, S.R.P. area, South
Carolina. 28 May 1985. (Coll. DGF #7496). (KE); (B) Sarsilatona serricauda Sars, 1901. Itatiba, Brazil. Figure
91 from Korovchinsky 1992.
21. Cladocera and Other Branchiopoda
877
2a (1a).
First antenna composed of two segments (one genus, Ilyocryptus, Fig. 44) ......................................................................Ilyocryptidae
2b.
First antenna with one segment ..........................................................................................................................................................3
3a(2a).
Second antenna (the swimming antenna) with one branch (Fig. 7) ...................................................................................Holopedidae
3b.
Second antenna with two branches ....................................................................................................................................................4
4a(3b).
First antennae fused with the rostrum to form two long and pointed tusklike structures (Figs. 41 and 42) ........................Bosminidae
4b.
First antennae not fused with the rostrum ..........................................................................................................................................5
5a(4b).
One branch of the second antenna with three segments, the other with four (look for a small basal segment, as in Fig. 1) ................6
5b.
Both branches of the second antenna with three segments (Suborder Radopoda) ...............................................................................7
6a(5a).
First antenna about as long as the width of the head; usually a tooth with two points (bifid) on the posterior angle of the postabdomen (near the claw), (Fig. 58)............................................................................................................................................Moinidae
6b.
First antenna less than half as long as the width of the head; teeth on postabdomen always single (Figs. 1A, 36 – 40)........Daphniidae
7a(5b).
First antenna is attached to the underside of the tip of the head, there is no rostrum (beak) (Figs. 44 – 57)........................Superfamily
Macrothricoidea
7b.
First antenna is attached to the ventral margin of the head, and is more or less covered with a rostrum (beak), (Figs. 13, 35)
..........................................................................................................................................................................................Chydoridae
8a(1b).
Body (thorax and abdomen) covered by the two valves (shells) of the carapace (Figs. 8 – 12)....................................................Sididae
8b.
Body not covered with a carapace (which is present in some forms as a saclike brood chamber attached to the back; legs are not
covered with the carapace (Figs. 60 – 64).....................................................................................Orders Onychopoda and Haplopoda
C. Taxonomic Key to Genera
of the Family Holopediidae
Hebert and Finston (1996) used results of allozyme
analysis to conclude that there are two species in North
America, Holopedium gibberum Zaddach, 1855 (Fig. 7)
and H. amazonicum Stingelin, 1904, confirming the
usage of Korovchinsky (1992). These are planktonic
animals, which often occur in soft and even acidic water.
H. gibberum is broadly distributed in the cool temperate
regions of North America, while H. amazonicum occurs
in the southern and eastern portions of the continent.
These species are remarkable in that each individual is
surrounded (out to about an additional body diameter)
by a mass of clear jelly attached to the carapace (easily
removed by rough handling). The jelly probably defends
Holopedium against an array of invertebrate predators.
Adult females are about 0.6 – 2.0 mm long.
D. Taxonomic Key to Genera of the Family Sididae
1a.
Posterior margin of carapace flared out laterally; a point on either posterior-lateral margin of the carapace; marine (Figs. 9B, C)
..................................................................................................................................................................................................Penilia
1b.
Posterior margin of carapace not flared laterally; postero-ventral corners of carapace rounded; freshwater .......................................2
2a(1b).
Swimming antenna appears to have three branches, the middle branch is an extension of the basal segment of the upper branch;
postabdominal claw with two basal spines (Fig. 11) .................................................................................................................Latona
2b.
Swimming antenna clearly has only two branches; postabdominal claw has 2 – 3 basal spines ...........................................................3
3a(2b).
Postabdominal claw with about 10 long teeth along the claw (“long” means longer than the width of the claw); a palearctic genus
............................................................................................................................................................................................Limnosida
3b.
Postabdominal claw with 2 – 4 long teeth near base............................................................................................................................4
4a(3b).
Postabdomen with a row of about 20 individual triangular spines along the margin (the anal margin); no clusters of spines on
postabdomen; postabdominal claw with three long teeth and an occasional short fourth tooth near the base of the claw; both
branches of swimming antennae with a total of 15 long swimming setae (Fig. 8) ..........................................................................Sida
4b.
Postabdomen with spines in clusters or patches, spines in rows are hairlike; swimming antennae with more than 15 long swimming
setae ...................................................................................................................................................................................................5
5a(4b).
Posterior margin of carapace with setae about 20 – 25% as long as the carapace ...............................................................................4
5b.
Posterior or ventral margin of carapace with setae shorter than 10% as long as the carapace ............................................................5
878
S. I. Dodson and D. G. Frey
6a(5a).
Swimming antennae branches with a total of 16 long swimming setae; postabdominal claw with two long basal spines (Fig. 11)
...........................................................................................................................................................................................Latonopsis
6b.
Swimming antennae branches with a total of 24 long swimming setae; postabdominal claw with three long basal spines (Fig. 12B)
...........................................................................................................................................................................................Sarsilatona
7a.(5b).
Swimming antennae with a total of 17 long swimming setae; postabdomen with rows of fine hairlike teeth; postabdominal claw
with three long teeth (the most basal tooth may be only about as long as the width of the claw at the base (Fig. 9) Diaphanosoma
7b.
Swimming antennae with a total of 20 long swimming setae, postabdomen with clusters of 4 – 6 spinelike teeth; postabdominal claw
with two long teeth, and a short basal tooth less than half as long as the width of the claw base (10) ................................Pseudosida
[Korovchinsky (1992) has recently revised the Sididae, including the genus Sarsilatona and retaining the changes made by Kořínek
(1981) for Diaphanosoma. Diaphanosoma and Penilia are planktonic and about 1.0 mm long; species in the other
genera tend to be larger (2 – 4 mm) and are associated with littoral vegetation.]
E. Taxonomic Key to Genera of the Family Chydoridae (North of Mexico)
1a.
Postabdomen strongly flattened, broad, with a single row of 80 – 150 marginal denticles, resulting in a saw like appearance; anus
distal; subfamily Eurycercinae (Fig. 13) ................................................................................................................................Eurycerus
[Genus is Holarctic except for isolated populations in southern Brazil – northern Argentina and in South Africa. At least five species in
North America, distributed among three subgenera: one species in the subgenus Eurycerus only within 300 km of the Gulf of Mexico
and the Atlantic Ocean as far north as the Pinelands of New Jersey (maximum length ca. 2 mm); two very common species in subgenus Bullatifrons, one in the northern tier of states and northward, the other in the south (length to 2.4 mm); and two species in the
subgenus Teretifrons in Newfoundland and the Arctic (length to 6 mm). See Frey (1971, 1975, 1978, 1982d) and Hann (1982).]
1b.
Postabdomen thicker, not so broad, and usually with two rows of marginal denticles, mostly fewer than 25 on each side; anus
proximal ........................................................................................................................................................................................... 2
2a(1b).
Proximal tip of mandible articulate where headshield and shell touch each other (Fig. 1C); usually two or three major headpores on
midline, which usually are connected by a double sclerotized ridge, resembling a channel (Fig. 1C); occasionally only one median
pore (Monospilus, Euryalona) or none (Notoalona); minor headpores lateral to these; free posterior margin of shell not greatly less
than maximum height of animal; typically one, occasionally zero, basal spine(s) on postabdominal claw (Fig. 1C); postabdomen
nearly always with lateral fascicles (Fig. 1C), sometimes without (Monospilus). ................................................Subfamily Aloninae 3
[In North America, Aloninae contains the genera Acroperus, Alona, Alonopsis, Bryospilus, Camptocerus, Euryalona,
Graptoleberis, Kurzia, Leydigia, Monospilus, Notoalona, Oxyurella, and Rhynchotalona. Olesen (1998) presents evidence that
Aloninae is a paraphyletic group.]
2b.
Proximal tip of mandible articulates in a special pocket on the headshield, some distance in a dorsal direction from the point of contact between the headshield and shell (Fig. 1D); two major headpores on midline (Fig. 1B, D), completely separated from one another and with no sclerotized ridge connecting them, occasionally only one pore (Dadaya), or only one pore in first instar and none
in later instars (Ephemeroporus); minor pores most commonly near midline between major pores; free posterior margin of shell
usually considerably less than maximum height of animal; typically two basal spines on postabdominal claw (Fig. 1D); articulated
lateral fascicles on postabdomen lacking, although Pseudochydorus has a row of fasciclelike groups of setae......................Subfamily
Chydorinae ......................................................................................................................................................................................15
[In North America, Chydorinae contains the genera Alonella, Anchistropus, Chydorus, Dadaya, Disparalona, Dunhevedia,
Ephemeroporus, Pleuroxus, and Pseudochydorus.]
3a(2a).
Only an ocellus present; no compound eye.........................................................................................................................................4
3b.
Compound eye and ocellus both present ............................................................................................................................................5
4a(3a).
Shells from previous instars firmly nested, so that instar number of specimen can be counted; headshield transversely trucate posteriorly; single median headpore (Fig. 14) .............................................................................................................................Monospilus
[Supposedly Monospilus contains just one species in world, mainly Holarctic and Ethiopian in distribution, with several records
from New Zealand as well. Length to 0.5 mm.]
4b.
Shells lost molting, so that no nesting of shells occurs; two separated median pores without sclerotized connecting ridge; minor
pores located far laterally; very short antennae (Fig. 15) ......................................................................................................Bryospilus
[Two species are known from New Zealand, Venezuela, and Puerto Rico, and hence this genus may possibly occur on north of
Mexico. Length to 0.35 mm. Found among wet, leafy hepatics of rain forests and cloud forests (Frey, 1980b, d).]
5a(3b).
Body strongly compressed from side to side, at times almost wafer thin; post-abdominal claw with a secondary spine midway along
concave margin, and usually with a comb of fine setules decreasing in length proximally between it and the basal spine (comb also
occurs in Euryalona) ..........................................................................................................................................................................6
5b.
Body not so strongly compressed; postabdominal claw having only a basal spine of variable length (except in Euryalona); sometimes none at all (one species of Leydigia) ..........................................................................................................................................9
21. Cladocera and Other Branchiopoda
879
6a(5a).
Body strongly keeled, and head also usually strongly keeled ..............................................................................................................7
6b.
Body weakly keeled, and head without a keel ....................................................................................................................................8
7a(6a).
Postabdomen narrow, elongate, tapered distally, with many (generally about 15 or more) margin denticles (Figs. 1C and 16)
......................................................................................................................................................................................Camptocercus
FIGURE 13 Eurycercus (Eurycercus) lamellatus (O.F. Mueller, 1776). Uppsala, Sweden. From Lilljeborg
(1901). FIGURE 14 Monospilus dispar Sars, 1862. Yddingesjon, Sweden. From Lilljeborg (1901). FIGURE
15 Bryospilus repens Frey, 1980. El Yunque, Puerto Rico. From Frey (1980b). FIGURE 16 Camptocercus
rectirostris Schoedler, 1862. Kristianstad, Sweden. From Lilljeborg (1901). FIGURE 17 Acroperus cf. harpae
(Baird, 1834). Eastern North America. From Birge (1918). FIGURE 18 Kurzia longirostris (Daday, 1898).
Tammanna wewa, Sri Lanka. From Rajapaksa (1986). FIGURE 19 Alonopsis elongata Sars, 1862. England.
From Norman and Brady (1867). FIGURE 20 Rhynchotalona falcata (Sars, 1861). Northern Michigan.
From Birge. (1893). FIGURE 21 Graptoleberis testudinaria (Fischer, 1851). Eastern North America. From
Birge (1918).
880
S. I. Dodson and D. G. Frey
[Nine species recognized in the world in 1971, with lengths from 0.7 to 1.26 mm. Except for C. oklahomensis, the species in North
America are completely unknown and undescribed, although certainly more than five species are present. C. rectirostris, C. liljeborgi,
and C. macrurus, which were described from Europe, do not occur in North America at all, even though reported by Brooks (1959).]
7b.
Postabdomen shorter and broader, parallel-sided, without any distinct marginal denticles (Fig. 17).....................................Acroperus
[Seven species in the world in 1971, with maximal lengths from 0.59 to 0.85 mm. Like Campocercus, the species in Acroperus are
not yet adequately described. There is marked conformity in body form and shape and in other morphologic characters, making it
very difficult to sort out the species satisfactorily.]
8a(6b).
Postabdomen rather slender, elongate, somewhat tapered distally; body high; shell widely open behind (Fig. 18) .....................Kurzia
[Five species in world and three in North America, K. latissima being widespread, and K. longirostris and K. polyspina occurring
only in the southernmost Gulf states. Length to 0.6 mm.]
8b.
Postabdomen broader, elongate, parallel-sided; shell with parallel striae sloping downward posteriorly, and with short longitudinal
scratch marks in between (Fig. 19)........................................................................................................................................Alonopsis
[Two species in world, the one in North America very similar to the one in Europe (see Kubersky, 1977). Occurs from northern
New England into the adjacent Canadian provinces and then into those farther East. Smirnov (1971) transferred the European
species to the genus Acroperus, which is not adequately justified. Maximum length: American 0.91 mm, European 0.85 mm.]
9a(5b).
Rostrum long, attenuate, recurved, greatly exceeding antennules in length (Fig. 20).....................................................Rhynchotalona
[Only one species listed in world, although there is a second species from North America not yet described. Postabdomen with 2 – 4
stout marginal denticles distally and with an almost continuous row of long, hairlike setae laterally. Length to 0.5 mm. A mud dweller.]
9b.
Rostrum rounded, barely or only slightly exceeding antennules in length.........................................................................................10
10a(9b).
Seen from above, rostrum very broad, semicircular, wider than body; shell and head strongly reticulated (Fig. 21) ........Graptoleberis
[Possibly only one species in world, although the taxonomy is not yet clear. Reported maximum length 0.7 mm. Ventral margin of
shell provided with a dense fringe of setae, and generally with two large, sharp, triangular teeth at posterior-ventral corner of shell;
postabdomen tapered distally, marginal denticles small, lateral fascicles minute; postabdominal claws small, with minute basal
spine. This species functions much like a snail, scraping food off the substrate as it moves along. See Fryer (1968).]
10b.
Rostrum more narrowly rounded; shell may be weakly reticulated, but head never is ......................................................................11
11a(10b).
Postabdomen elongate and rather narrow; with marginal denticles well developed ..........................................................................12
11b.
Postabdomen much less elongate (except in Leydigia); marginal denticles usually well developed, but in some species greatly
reduced ............................................................................................................................................................................................13
12a(11a).
Dorsal margin of postabdomen straight or slightly convex; marginal denticles very short proximally, increasing in length distally to
three very long curved denticles; four median headpores (middle one is divided into two), usually completely separated from one
another, or with middle pores joined by a sclerotized ridge (Figs. 21. 1C and 22) ................................................................Oxyurella
[Basal spine of postabdominal claw long, slender, attached some distance from base of claw; head and shell generally have a yellowish, hyaline appearance. Five species claimed to occur in the world in 1971. Two species in North America, one being confined to
the Gulf states, the other more widely distributed in the East. The common North American species is now separated from the
formerly cognate species in Eurasia. See Michael and Frey (1983).]
12b.
Dorsal margin of postabdomen distinctly concave; marginal denticles increase in length distal, but never reach large size as in
Oxyurella; one median headpore, sometimes completely absent (Fig. 23) ............................................................................Euryalona
[A tropical to subtropical genus. Three species in world, the one in America north of Mexico originally described from Sri Lanka.
Head high; rostrum broadly rounded, with tip scarcely reaching halfway to ventral margin of shell; comb of setae on proximal half
of concave margin of postabdominal claw, increasing in length distally; stout spine on inner distal lobe of trunklimb I, with coarse,
rounded tubercles on concave edge. See Daday (1898) and Rajapaksa (1986).]
13a(11b).
Postabdomen large, broad, flattened, almost semicircular; armed laterally with long, spinelike setae in groups, those of distal groups
projecting far beyond margin of postabdomen; margin of postabdomen with short, fine spinules; three median headpores close together, and with minor pores close-in laterally (Fig. 24) ..........................................................................................................Leydigia
[Free posterior margin of shell very long; ocellus as large as compound eye or larger. Twelve species claimed for world in 1971, with
maximal lengths from 0.8 to 1.15 mm. Two species are widely distributed in United States, and one tropical species barely gets into
Florida.]
13b.
Postabdomen smaller, and with shorter setae in lateral fascicles; headpores variable ........................................................................14
14a(13b).
Shell strongly sculptured, with longitudinal striae in posterior part and about five vertical striae anteriorly, roughly parallel to the
anterior margin; dorsal edge of postabdomen minutely serrate; about 14 fascicles laterally; no median headpores, but instead there
are two comma-shaped thickenings, which presumably contain the minor pores (Fig. 25) ..................................................Notoalona
[A genus containing two tropical to subtropical species, only one of which gets into the United States. The headpore configuration is
unique among the Chydoridae. Length to 0.44 mm. See Rajapaksa and Fernando (1987a).]
21. Cladocera and Other Branchiopoda
14b.
881
Shell usually not sculptured at all, or if so, then rather weakly; postabdomen usually with well-developed marginal denticles and
lateral fascicles; two or three median headpores, most commonly united by a sclerotized ridge, although sometimes completely
separated (Fig.26) .......................................................................................................................................................................Alona
[This is the largest genus in the Chydoridae, with 52 species claimed in the world in 1971 and with more than 20 species in the
United States and Canada. Alona intuitively consists of a number of genera, but these have not yet been defined satisfactorily.
FIGURE 22
Oxyruella brevicaudis Michael and Frey, 1983. Skater’s Pond, Monros Co., Indiana. From
Michael and Frey (1983). FIGURE 23 Euryalona orientalis (Daday, 1898). Ipiranga, Brazil. From Rajapaksa
(1986). FIGURE 24 Leydigia acanthocercoides (Fischer, 1854). Uplandia, Sweden. From Lilljeborg (1901).
FIGURE 25 Notoalona freyi Rajapaksa and Fernando, 1987. Homer Lake, Leon Co., Florida. From
Rajapaksa and Fernando (1987a). FIGURE 26 Alona bicolor Frey, 1965. Goodwil Pond, Barnstable Co.,
From Massachusetts. From Frey (1965). FIGURE 27 Dadaya macrops (Daday, 1898). Polonnaruwa, Sri
Lanka. From Rajapaksa and Fernando (1982). FIGURE 28 Ephermeroporus acanthodes Frey, 1987.
Audobon Park, New Orleans. From Frey (1982b). FIGURE 29 Dunhevedia americana Rajapaksa and
Fernando 1987. Glades Co., Florida. From Rajapaksa and Fernando (1987b).
882
S. I. Dodson and D. G. Frey
Smirnov (1971) placed the species having just two median headpores in a separate genus, Biapertura. Three of the species in North
America, currently named A. affinis, A. intermedia, and A. verrucosa, belong here. Smirnov and Timms (1983) found a much
greater proportion of two-pored species in Australia, which do not seem to be closely enough related to be included in the same
genus. Length of species in America north of Mexico: two-pored species to 1.05 mm; three-pored species 0.4 – 0.9 mm.]
15a(2b).
Headshield not elongated posteriorly; in two-pored species, postpore distance usually considerably less than interpore distance.....16
[Includes the genera Alonella, Dadaya, Disparalona, Dunhevedia, and Ephemeroporus.]
15b.
Headshield much elongated posteriorly, extending well beyond the heart to the middle of the back; postpore distance usually
greater than interpore distance, sometimes by several-fold (Fig. 1D) ................................................................................................19
[Includes the genera Anchistropus, Chydorus, Pleuroxus, and Pseudochydorus.]
16a(15a).
Only one median headpore, or none ................................................................................................................................................17
16b.
Two median headpores, well separated and not connected by a sclerotized ridge (Fig. 1D) ..............................................................18
17a(16a).
One median headpore in all instars (Fig 27); ocellus larger than eye ........................................................................................Dadaya
[A single pantropical species occurring sparingly in the Gulf states. Color dark brown. Length to 0.41 mm.]
17b.
One median headpore only in first instar; none in later instars (Fig 28); ocellus smaller than eye ................................Ephemeroporus
[Postabdomen with long proximal and distal marginal denticles, and with shorter ones in between; labral keel large, with 1 – 4 teeth
on anterior margin. A tropical to subtropical genus with many species, few of which have been described to date. See Frey (1982b).]
18a(16b).
Postabdomen unique among chydorids, consisting of a much expanded postanal portion, having a series of short spines along dorsal margin and many small clusters of spinules laterally (Fig. 29) ......................................................................................Dunhevedia
18b.
Postabdomen of more typical chydorid structure, with pre- and postanal angles and a well-developed series of postanal marginal
denticles (Figs. 30 and 31) ............................................................................................................................Alonella and Disparalona
[Four species claimed for world, two in the United States, one of which is strictly southern. Length for 0.46 – 0.8 mm.]
[The taxon Disparalona rostrata of Europe was formerly in the genus Alonella. Fryer (1986) removed it from this genus because it
had a long “sweeping” seta on trunklimb III, which the other species of Alonella were claimed not to posses. Fryer (1971) subsequently transferred Alonella acutirostris to Disparalona, and Smirnov (1971) did the same for Alonella dadayi. Michael and Frey
(1984) closely examined a number of species of Alonella and of Disparalona and concluded that the sweeper seta is not an additional seta in a few species but rather is the proximal member of the gnathobasic filter comb, and that species still in Alonella have
the same seta variably developed and not always distinctly separable from the Disparalona species. I know of no characters that
will clearly separate the two groups of species, suggesting that the validity of Disparalona is questionable. Disparalona presently
contains the species D. acutirostris, D. dadayi, D. leei, all occurring in North America, and Alonella the species A. excisa, A.
exigua, A. hamulata, A. nana, and A. pulchella, out of a world total of 12. Maximal lengths of the North American taxa in Disparalona are 0.45 – 0.5 mm, and in Alonella, 0.25- 0.6 (Michael and Frey 1984, Hann and Chengalth 1981).]
19a(15b).
Body elongate, somewhat flattened, usually with one or more teeth at the posterior-ventral angle of shell (Figs. 1D and 32)
.............................................................................................................................................................................................Pleuroxus
[Fifteen species claimed in world, with maximal lengths from 0.4 – 0.8 mm. Seven species in America north of Mexico, of which P.
procurvus and P. denticulatus are most frequent and most abundant (Frey 1988c). Lengths from 0.5 – 0.8 mm.]
19b.
Body globular, nearly round, seldom with any teeth at posteor-ventral angle ...................................................................................20
20a(19b).
Ventral margin of shell anteriorly with hooklike development containing a groove, in which a stout seta with teeth on trunk limb I
operates; postabdomen with a cluster of long, slender setae at distal angle (Fig. 33)........................................................Anchistropus
[Surface of shell and head reticulated, with meshes having wavy edges. Three species in world, at least two of which are parasitic on
Hydra. Lengths 0.35 – 0.46 mm, the North American species being the smallest. See Hyman (1926) and Smirnov (1985).]
20b.
Ventral margin of shell without such a hook; marginal setae of shell arise submarginally posteriorly, forming a distinct duplicature
(except in Chydorus piger) ...............................................................................................................................................................21
21a(20b).
Postabdomen elongate, parallel-sided; postanal portion with about 15 slender marginal denticles; lateral surface with a row of
fasciclelike clusters of setae (Fig. 34) ...........................................................................................................................Pseudochydorus
[Pseudochydorus presently contains a single species, distributed over much of the world, except possibly in South America. Length
to 0.8 mm. It is a scavenger. See Fryer (1968).]
21b.
Postabdomen shorter, with a smaller number of marginal denticles, and laterally with crescentic clusters of short spinules, which do
not resemble fascicles (Fig. 35) ..............................................................................................................................................Chydorus
[A highly complex genus, in which the species are difficult to separate, because they conform so closely to a common morphotype.
At least ten species have been found in America north of Mexico. Smirnov (1971) recognized 18 species and many subspecies in the
world, ranging in maximum length from 0.32 – 1 mm. The species occurring in North America range in length from 0.41 to
0.78 mm. The type of species of the family, Chydorus sphaericus (sens. str.), quite possibly does not occur in North America at all.
See Frey (1980a, 1982a, c, 1987b).]
21. Cladocera and Other Branchiopoda
883
FIGURE 30 Alonella pulchella Herrick 1884. Kremer Lake, Minnesota. From Hann and Chengalath (1981).
FIGURE 31 Disparalona leei (Chien 1970) Griffey Lake, Monroe Co., Indiana. From Michael and Frey
(1984). FIGURE 32 Pleuroxus procurvus Birge, 1878. Lake Mendota, Dane Co., Wisconsin. 11 September
1987. (KE) FIGURE 33 Anchistropus minor Birge, 1893. Eastern North America. From Birge (1918).
FIGURE 34 Pseudochydorus globosus (Baird, 1843). River Sutka, USSR. From Smirnov (1971). FIGURE 35
Chydorus brevlabris Frey, 1980. Salmon Lake, Montana. From Frey (1980a).
F. Taxonomic Key to Genera of the Family Daphniidae and Moinidae
Fryer (1991) presents a complete and detailed analysis of the functional morphology of genera of the Daphniidae.
1a.
First antenna about as long as the width of the head; usually a tooth with two points on the posterior angle of the Bostabdomen
(near the claw), (Fig. 58) ........................................................................................................................................Family Moinidae 7
1b.
First antenna less than half as long as the width of the head; teeth on postabdomen always single (Figs. 1A, 36 – 40).
...........................................................................................................................................................................Family Daphniidae 2
2a.
Ventral margin of carapace rounded and not pigmented ....................................................................................................................3
2b.
Ventral margin of carapace straight and usually black........................................................................................................................6
884
S. I. Dodson and D. G. Frey
FIGURE 36
Ceridodaphnia cf. quadrangula (O.F. Mueller, 1785). Stewart Lake, Dane Co., Wisconsin. 5
September 1973. (KE) FIGURE 37 Simocephalus (A) S. vetulus Schoedler, 1858. Gardener Pond, UW
Arboretum, Dane Co., Wisconsin. 14 May 1987. (KE) (B) S. serrulatus (Koch, 1841). Coliseum Pond,
Madison, Dane Co., Wisconsin. 15 May 1987. FIGURE 38 Daphniopsis ephemeralis Schwartz and Hebert
1985. Baseline Pond, Ontario. 19 April 1988. (Coll. P.D.N. Hebert). (KE) FIGURE 39 Megafenestra nasuta
(Birge, 1879). Dorsal view. Elk Island National Park, Alberta. 5 August 1978. (KE). Redrawn from Dumont
and Pensaert (1983). FIGURE 40 Scapholeberis rammneri Dumont and Pensaert, 1983. Ojibwa, Ontario. 19
April 1988. (Coll. P.D.N. Hebert). (KE)
3a(2a).
Adults with a tail spine, at least four times as long as broad and pointed (Figs. 1, 2, 4, 5)......................................................Daphnia
3b.
Adults lack a tail spine, or if one is present, it is less than four times as long as broad and rounded ..................................................4
4a(3b).
Fourth (distal) segment longer than the third segment on the four-segment branch of the second (swimming) antenna (Fig.38)
.........................................................................................................................................................................................Daphniopsis
[One species, D. ephemeralis in North America (Schwartz and Herbert 1987c); the genus is reviewed by Hann (1986).]
4b.
Fourth segment shorter than the third segment on the four-segment branch of the second antenna ...................................................5
21. Cladocera and Other Branchiopoda
5a(4b).
885
The second segment of the four-segment branch of the second antenna with a spine, the spine about 1 / 4 as long as the second
segment (Fig 37) .............................................................................................................................................................Simocephalus
[Allozyme studies of Simocephalus suggest there are at least five species in North America, two more than are recognized using
morphological characters (Hann and Herbert 1986). Orlova-Bienkowskaja (1998) recognized at least eight North American
species.]
5b.
The second segment of the four-segment branch of the second antenna without an apical spine (Fig. 39) .......................Ceriodaphnia
[Species of Ceriodaphnia were discussed by Brandlova et al. (1972), but are still poorly know and in need of taxonomic attention.]
6a(2b).
Dorsum of headshield with a median oval plate about 20 m in diameter, marked by slightly raised edges (Fig 39) .......Megafenestra
6b.
Dorsum of headshield without such a plate (Fig 40) ......................................................................................................Scapholeberis
[With the exception of Megafenestra and Scapholeberis (Dumont and Pensaert 1983), and perhaps Daphniopsis (Hann, 1986;
Schwartz and Hebert, 1987c), the daphniid genera are in need of revision. Daphnia are being carefully studied by Hebert (Hebert,
1995; Colbourne et al., 1997), who recognizes about 34 species in North America. All of the daphniid genera tend toward the planktonic lifestyle. Megafenestra and Scapholeberis are the least planktonic; they are often found cruising along upside-down, at the surface of small pools. Their carapaces have flattened and turned-in margins (Dumont and Pensaert 1983), which may allow the carapace to serve as a suction cup, as in some chydorids. Simocephalus spends much of its time stuck to aquatic plants, using sticky
mucous produced at the back of the neck. Daphniopsis has an extremely restricted habitat; it has been found in North America only
in shallow temporary pools in maple forests. Daphnia and Ceriodaphnia are often restricted to open water of lakes and ponds. Scapholeberis and Ceriodaphnia adults are small, reaching about 1 mm, while the other genera produce adults in the range of 1 to 3 mm.]
7a.(1a).
Branch of second antenna with four segments appears to have four terminal setae; ocellus (a small black spot just back of the eve)
absent in North American species (Fig. 58) ................................................................................................................................Moina
7b.
Branch of second antenna with four segments has three setae and a short spine; an ocellus is present (Fig. 59).............Moinodaphnia
[These two genera were monographed by Goulden (1968). The species are typically planktonic, but the majority are restricted to
small temporary ponds, saline, or alkaline lakes. The habitat is often ephemeral, turbid, and warm. Adult females reach 1 – 2 mm,
with one Moina species that is only about 0.5 mm long.]
G. Taxonomic Key to Genera of the Family Bosminidae
1a.
First antennae attached separately to the head, immobile in female, usually nearly vertical and usually curving somewhat posteriorly
(Fig. 42A) ...............................................................................................................................................................................Bosmina
(with four subgenera) .........................................................................................................................................................................2
1b.
First antennae fused together in basal half, separated and curved outward in distal half (resembling a mermaid), firmly attached to
head (Fig. 41) ...................................................................................................................................................................Bosminopsis
2a(1a).
Lateral headpore located within five pore diameters of the edge of the headshield, just dorsal to the base of the second antenna and
at the bottom of the reticulated part of the headshield (Figs. 42C, 43D); teeth of distal pecten (comb of spines) on postabdominal
claw as wide as long and resemble an equilateral triangle ..................................................................................................................3
2b.
Lateral headpore located more than 20 pore diameters from the edge of the headshield, near the point of articulation of the
mandibles, in the reticulated region of the headshield; teeth of distal pecten much longer than wide and resemble needles ................4
3a.(2a).
Lateral head pore next to the basal striation at edge of the headshield (Fig. 42C); teeth of the proximal pecten of the postabdominal
claw are hairlike, only the largest tooth is perceptibly triangular ........................................................Bosmina (Bosmina) longirostris
3b.
Lateral head pore between the two branches of the basal striation; teeth of the proximal pecten of the postabdominal claw are narrow triangles ...................................................................................................................................................Bosmina (Sinobosmina)
4a(2b).
Mucro (tail spine) absent in juvenile female (Fig. 43A), or if present, with minute incisions present only along the ventral margin;
incisions may be lacking in the adult female; mostly in northern locations ........................................................Bosmina (Eubosmina)
4b.
Mucro present in juvenile female, with minute incisions or teeth present only on the dorsal margin (Fig. 42D); mostly in southern
locations ......................................................................................................................................................... Bosmina (Neobosmina)
[There are just two genera in the family Bosminidae. The key to the subgenera of Bosmina is based on DeMelo and Hebert, (1994).
The subgenus B. Bosmina contains the single species B. (B.) longirostris, which until DeMelo and Hebert (1994) had been considered to be very common. However, their revision indicated that B. (B.) longirostris is rare, and that instead most records probably
should be for the subgenus Sinobosmina, either B. (S.). leideri or B. (S.) freyi. There are currently four North American species in
the subgenus Eubosmina and three North American species of Neobosmina. The subgenera are separated using the shape of the
carapace, spines on the postabdominal claw, and the position of the lateral head pores on the head shield (Fig. 42E – G). Bosminids
are usually planktonic, with Bosminopsis the least so. Adult bosminids range in size from about 0.3 – 0.6 mm long. Because of their
small size, bosminids often are abundant when fish predation is intense.]
FIGURE 41 Bosminopsis deitersi Richard 1895. Singletary Lake, North Carolina. 8 August 1948. (coll. DGF
#236. (KE) FIGURE 42 (A) Bosmina (Bosmina) longirostris (O.F. Mueller, 1776). Lake Mendota, Dane Co.,
Wisconsin. 11 September 1987. (KE); (B) Postabdomen of A (KE); (C) Headshield of A, (KE); (D) Headshield of
20.43A. (KE); (E) Head pore position typical of Bosmina (Sinobosmina), (DeMelo and Hebert 1994); (F) Head
pore position typical of Bosmina (Bosmina), (DeMelo and Hebert 1994); (G) Head pore position typical of
Eubosmina (Bosmina) and Bosmina (Neobosmina), (DeMelo and Hebert 1994) FIGURE 43 (A) Bosmina
(Eubosmina) coregoni (Biard, 1850). Lake Waubesa, Dane Co., Wisconsin, 23 July 1988; (B) Postabdomen of
A. (KE); (C) Bosmina (Neobosmina) tubicen Brehm, 1953. Presa Presidente Calles. Aquascalientes, Mexico. 8
October 1980. (KE); (D) Bosmina (Sinobosmina) fatalis Burchardt 1924. Redrawn from Kořínek (1971). (KE)
886
21. Cladocera and Other Branchiopoda
H. Key to the Genera of Superfamily Macrothricoidea
This key makes extensive use of setae on the second
antenna (AII, or swimming antenna). A seta is longer
than the segment it sits on, and it resembles a feather,
having fine setules. A spine is usually shorter than the
segment it sits on, and lacks the setules. The AII has two
branches: the dorsal branch with four segments (look for
a small segment at the base of one of the branches), and
887
a ventral branch with three segments. For best results,
dissect at least one AII off the specimen, and mount it so
the branches and setae are spread out. This key is for
adult females only, and animals are arranged to facilitate
ease of identification, not to reflect phylogenetic relationships. For further details on morphology, see Smirnov
(1992), Dumont and Silva-Briano (1998) and SilvaBriano (1998). For a careful morphological analysis of
the macrothricids (and related taxa) see Olesen (1998).
1a.
Carapace and head covered with fine hairlike setae (Cactus is South American, Neothrix is Australian; Family Neothricidae) (Fig.
56) .......................................................................................................................................................................Cactus and Neothrix
1b.
Carapace not covered with hairlike setae ...........................................................................................................................................2
2a.(1).
AII dorsal (four-segmented) branch with no setae except for the three terminal setae.........................................................................3
2b.
At least one of the subterminal segments of the AII dorsal branch with a seta (Family Macrothricidae, in part) ................................8
3a.(2).
Postabdominal claw with two basal spines, each longer than the width of the claw (Family Ophryoxidae) .......................................4
3b.
Postabdominal claw without basal spines, or with small spines much shorter than the width of the claw ..........................................5
4a.(3).
The three-segmented branch of AII with a seta at the distal end of each of the two subapical segments (Family Ophryoxidae) (Fig.
45)......................................................................................................................................................................................Ophryoxus
4b.
The three-segmented branch of AII with no seta on the two subapical segments (Fig. 46) ..............................................Parophryoxus
5a.(3).
The posterio-ventral corner of the carapace with setae about half as long as the carapace height (Family Acantholeberidae) (Fig. 47)
.....................................................................................................................................................................................Acantholeberis
5b.
The posteroventral corner of the carapace with setae less than 0.1 times as long as the height of the carapace (Family Macrothricidae, in part)........................................................................................................................................................................................6
6a.(5).
Dorsal margin of the carapace with a small dorsal protrusion (about the size of the compound eye) .............................Onchobunops
6b.
Dorsal margin of the carapace without a rounded protrusion ...........................................................................................................7
7a.(6).
Dorsal margin of carapace with a backward-pointing tooth, dorsal margin of first antenna with teeth about 0.3 times as long as the
width of the antenna a Palearctic genus) (Fig. 48) ...........................................................................................................Drepanothrix
7b.
Dorsal margin of carapace without a backward-pointing tooth, margins of first antenna smooth or with small teeth less than 0.1
times as long as the width of the antenna (Fig. 53) ...................................................................................................................Bunops
8a.(2).
Segments 2 and 3 (counting from the base) of the four-segmented branch of AII with setae ...............................................................9
8b.
Only segment 3 of the four-segmented branch of AII with a seta......................................................................................................11
9a.(8).
Ventral margin of the carapace with flattened spines (“lanceolate”) that are about twice as long as wide (Fig. 50) .............Lathonura
9b.
Ventral margin of the carapace with cylindrical spines that are several times as long as wide...........................................................10
10a.(9).
Postabdominal claw with a basal spine that is longer than the width of the claw (an Australasian genus) (Fig. 57).........Pseudomoina
10b.
Postabdominal claw without a basal spine (a circum-tropical genus) (Fig. 52) ......................................................................Guernella
11a.(8).
Postabdomenal claw with a basal spine that is longer than the width of the claw (this character shown in the drawing in SilvaBriano, 1998, but not in Smirnov, 1992); a single large spine near the dorsal margin of the opening of the anus, the spine nearly as
long as the width of the opening (a circum-tropical species) (Fig. 51).................................................................................Grimaldina
11b.
Postabdominal claw without a basal spine, and anus opening without long spines ..........................................................................12
12a.(11).
Postabdomen with large saw teeth along the margin; first antenna with a basal seta and four spinelike setae, all of which are longer
than the width of the antenna (Fig. 49) .............................................................................................................................Streblocerus
12b..
Postabdomen with fine hair-like teeth along the margin; first antenna with a basal seta and some other arrangement of setae toward
the tip...............................................................................................................................................................................................13
13a.(12).
First antena with terminal seta that are less than 1 / 4 as long as the antenna, the setae are nearly the same length and thickness, the
first antenna is cylindrical, not inflated toward tip (Fig. 55) .................................................................................................Wlassicsia
13b.
First antenna with at least two of the terminal setae that are at least 1 / 3 as long as the antenna, two of the terminal setae are at
least 50% longer and thicker than the other setae; the tip of the first antenna may be wider at the tip than the base (Fig. 54)
...........................................................................................................................................................................................Macrothrix
888
S. I. Dodson and D. G. Frey
FIGURE 44
Ilyocryptus sordidus (Liévin 1848). English lake District. From Fryer (1974). FIGURE 45
Ofryoxus gracilis Sars, 1861. English Lake District. From Fryer (1974). FIGURE 46 Paraphryoxus tubulatus
Doolittle, 1909. Anonymous Pond, Maine. From Doolittle (1911). FIGURE 47 Acnatholeberiscurvirostris
(O.F. Mueller, 1776). English Lake District. From Fryer (1974). FIGURE 48 Drepanothrix dentata (Eurén,
1861). English Lake District. From Fryer (1974). FIGURE 49 Streblocerus serricaudatus (Fischer 1849).
English Lake District. From Fryer (1974). FIGURE 50 Lathonura rectirostris (O. F. Mueller 1776). Near
Leningrad, USSR. From Smirnov, after Sergeyev (1971).
21. Cladocera and Other Branchiopoda
889
FIGURE 51
Grimaldina brazzai Richard, 1892. Louisiana. From Birge (1918). FIGURE 52 Guernella
raphaelis Richard, 1892. Mayumba, Gabon. From Richard (1892). FIGURE 53 Bunops acutifrons Birge,
1893. Minocqua, Wisconsin. From Merrill (1893). FIGURE 54 Macrothrix rosea (Jurine, 1820). Hook Lake,
Dane Co., Wisconsin. 25 October 1972. (KE) FIGURE 55 Wlassicsia kinistinensis Birge, 1910. Kinistino,
Manitoba. From Birge (1910). FIGURE 56 Neothrix armata Gurney, 1927. Fig. 491 from Smirnov (1992).
FIGURE 57 Pseudomoina lemnae (King, 1853). Fig. 475 from Smirnov (1992).
I. Key to the Orders Onychopoda and Haplopoda
See Rivier (1998) for details of these predatory cladocerans.
1a.
Head cylindrical, more than twice as long as wide, swimming antennae with 36 setae (both branches) (Order Haplopoda, Fig. 60)
............................................................................................................................................................................................Leptodora
[Leptodora kindtii (Fig. 60) is the sole species in this group. The adult females are up to 18 mm long. Despite their large size, the
animals are so transparent as to be nearly invisible. They are common in the open water of large lakes, and occasionally are found
in ponds. L. kindtii are voracious predators on smaller zooplankton.]
890
S. I. Dodson and D. G. Frey
FIGURE 58 Moina wierzejskii Richard, 1895. 45 km NE of Las Cruces, New Mexico. 6 July 1984 (KE)
FIGURE 59 Moinodaphnia macleayii (King, 1853). Roadside canal, Highlands Co., Florida. 20 July 1960.
(Coll. DGF #96). (KE) FIGURE 60 Leptodora kindti (Focke, 1844). Lake Michigan, USA. From Balcer et al.
(1984). Drawn by Nancy Korda. FIGURE 61 Bythotrephes cederstroemii Schodler, 1877. Summer female
from Lake Karesuando in Norrbotten. From Lilljeborg (1901). Redrawn by KE. FIGURE 62 Polyphemis
pediculus (Linnee, 1761). Lake Michigan, USA. From Balcer et al. (1984). Drawn by Nancy Korda. FIGURE
63 Evadne nordmanni (LovÇn, 1836). Summer female from the sea near Dalaro. From Lilljeborg (1901).
Redrawn by Cheryl Hughes. FIGURE 64 Podon leuckartii Sars, 1862. Summer female from the sea near
Bergen, Norway. From Lilljebory (1901). Redrawn by Cheryl Hughes.
21. Cladocera and Other Branchiopoda
891
1b.
Head not obviously cylindrical, swimming antennae with 15 or fewer setae (both branches) (Order Onychopoda) ...........................2
2a (1b).
Adult female total length 1 – 2 mm, abdominal process shorter than the body....................................................................................3
2a.
Adult female total length greater than 2 mm, abdominal process longer than the body ( Fig. 61, Family Cercopagidae)....................5
3a (2a).
Abdominal process slender and about half as long as the body, freshwater species in North America (Family Polyphemidae, Fig. 62)
Polyphemus
[The single species in North America (Polyphemus pediculus) is often found along the margins of small bog lakes.]
3b.
Abdominal process blunt and shorter than half of the body length, estuarine or marine species along the coasts of North America
(Family Podonidae) ............................................................................................................................................................................4
4a (3b).
Body ends in a short extension of the abdomen, about twice as long as broad (Fig. 63) ...........................................................Evadne
4b.
Body ends in a triangular point, but without a distinct abdominal process (Fig. 64) ..................................................................Podon
5a (2a).
Swimming antennae with 15 setae (both branches) (Fig. 61) ...........................................................................................Bythotrephes
5b.
Swimming antennae with 14 setae (both branches) .............................................................................................................Cercopagis
[Bythotrephes longimanus and Cercopagis pengoi are invaders of the Lawrentian Great Lakes, having come from Europe, probably in
ballast water. See the Sea Grant web page at http: / /www.ansc.purdue.edu / sgnis / home.htm B. longimanus (probably from a Polish
harbor) was first seen in Lake Huron in 1984 and C. pengoi (from the Caspian-Black Sea region) appeared in Lake Ontario in 1998. ]
OTHER BRANCHIOPODS
XIII. ANATOMY AND PHYSIOLOGY
OF THE OTHER BRANCHIOPODS
A. General External and Internal Anatomic Features
Martin (1992) provides a masterful and detailed
description of branchiopod morphology, with many
drawings and photographs. The non-cladoceran branchiopods are composed of four extant orders (Table I)
They differ from cladocerans in a number of ways, including having more pairs of legs and paired compound eyes. The four orders are probably not closely
related and are morphologically diverse (McLaughlin,
1980; Fryer, 1987).
Members of the first order, Anostraca (fairy
shrimps), swim with their legs up (Fig. 65), have a pair
of large compound eyes, and possess no shell-like
covering. Anostracan morphology is described in detail
by Linder (1941). Adults are 7 – 100 mm long and have
11, 17, or 19 pairs of thoracic legs along their rather
delicate, transparent, and elongate body. The legs are
all anterior to two fused genital segments (Fig. 65);
hence, they can be called thoracic legs. There are no
abdominal legs, but rather a pair of terminal lobes
(cercopods, Fig. 67) on the last abdominal segment
(telson). The genital segments contain the gonads
(which extend into abdominal and thoracic segments).
Males have a pair of ventral penes (Figs. 70C and 71).
Females possess a medical brood pouch (Fig. 70B).
Within the brood pouch are lateral oviducal pouches, a
single median ovisac, and shell glands. Males have
large and sometimes complex second antennae
(Fig. 69A) and sometimes outgrowths from the
head (Fig. 66C) or antennae (Fig. 70A). The second
antennae are used for grasping females (Fig. 73B) but
not for swimming (as in cladocerans and cochostracans).
Laevicaudata and Spinicaudata (the two conchostracan orders of clam shrimps, Figs. 75 and 76) swim
with their legs down or forward, have a pair of closeset compound eyes (separated, except in Cyclestheria
where they are fused), and a carapace that wraps
around the entire body (Figs. 75 and 76). Adults are
2 – 16 mm long and have 10 – 32 pairs of thoracic legs
covered by the transparent to brown carapace
(Fig. 76). In species of Spinicaudata, the carapace
closely resembles a clam shell, even having growth lines
and umbo (Fig. 81). In Laevicaudata, the carapace is
more globular, and only one Siberian) species has even
a single growth line. The branched armlike appendages
that can be extended from the carapace are the second
antennae (Fig. 81); they are used for swimming, as in
the cladocera.
Notostraca, or tadpole shrimps, are sometimes
confused with trilobites by the unwary. These crustaceans swim with their legs down, have a pair of dorsal compound eyes, and possess a broad and flat carapace that covers the head and thorax (Fig. 82). Adults
are 10 – 58 mm long and have 35 – 70 pairs of trunk
legs partially hidden by their dark green or grown carapace. Immature notostracans have a transparent carapace that is folded along the dorsal midline and encloses the body and legs, making them appear
somewhat like a conchostracan. Long filaments (endites) protruding from either side of the carapace are
attached to the first pair of thoracic legs (Fig. 82).
892
S. I. Dodson and D. G. Frey
FIGURE 65
Polyartemiella hazeni (Murdoch, 1874). Pond #14, 18 July 1973, Point Barrow, Alaska. 18 July
1973. (A) female; (B) male; (C) head of male. (KE) FIGURE 66 Thamnocephalus platyrus Packard, 1879.
Pond near Ringold, McPherson Co., Nebraska. 28 June 1972. (Coll. S. Cooper). (KE) (A) Female abdomen
from side; (B) male abdomen from below; (C) male head; (D) detail of spines on frontal process. FIGURE 67
Branchinella sublettei Sissom, 1976. (Coll. D. Belk #717). (A) Head of male; (B) male abdomen. (KE) FIGURE
68 Artemia franciscana (Kellogg 1906). Male head. Alkali lake, Owen’s Valley, California. 2 July 1974. (KE)
FIGURE 69 Streptocephalus texanus Packard, 1971. Texas. (Coll. F. Wiman). (A) Head of male, second antenna unrolled; (B) Head of male, second antenna coiled. (KE)
21. Cladocera and Other Branchiopoda
Endites of the second pair of legs are less filamentous,
and the succeeding legs lack filaments (Fryer 1988).
The first pair of legs is used in swimming. Phyllopods
posterior to the first pair are used for swimming and
also walking, digging, and food handling (D. Belk, personal communication; Fryer, 1988).
Eggs are carried in brood pouches in the specialized eleventh pair of legs. The anterior lid of the concave brood pouch is derived from the exopodite of the
leg and the bottom from the subapical lobe of the leg
(Fryer, 1988). The eleventh pair of thoracic legs of
males resembles the tenth and twelfth pairs. The gross
morphological difference between males and females is
that males lack the brood pouch (and ovaries) and tend
to be slightly larger than the hermaphroditic “females”
(Beaton and Hebert, 1988).
Maleless populations are usually (Akita, 1971;
Beaton and Hebert, 1988; Longhurst, 1954; Fryer,
1988), but not always (Zaffagnini and Trentini, 1980)
composed of hermaphrodites. Zaffagini and Trentini
(1980) present evidence that reproduction in maleless
populations is by automictic parthenogenesis (i.e., selffertilizing hermaphrodites), not autogamic parthenogenesis as previously suggested (Longhurst, 1954).
Beaton and Hebert (1988) found no electrophoretic
variation in Lepidurus arcticus and were not able to
clarify the role of males in reproduction.
The notostracan trunk is not properly divided into
thorax and abdomen, because the legs continue past
the genital opening on the eleventh segment and there
are many more legs than “segments.” The legs gradually decrease in size toward the tail. The body ends in a
pair of long thin cercopods (Fig. 82); and in Lepidurus,
a posterior extension of the telson (supra-anal plate,
Fig. 82A).
B. Relevant Physiological Information
Branchiopods of the different orders are similar in
regard to their physiological adaptations. Thus, the discussion of cladoceran physiology applies in general to
the noncladoceran orders as well. Artemia is a model
animal for physiologists (e.g., Sorgeloos et al., 1987).
Conchostracans, notostracans, and anostracans
probably depend more on gills for respiration than do
the cladoceran orders (Eriksen and Brown, 1980a).
Notostracans and anostracans show the usual correlation between surface area and the rate of oxygen
consumption, although in conchostracans, small juveniles have an unusually high VO2 (rate of oxygen uptake), perhaps due to the well-developed gills on the
trunk legs.
Osmoregulation (maintenance of proper salt concentration in the blood and tissues) is important to
893
these animals, which often live in conditions of extreme
or variable salinity. The general mechanisms of osmoregulation are probably similar in all branchiopods
(Peters, 1987), but there are many variations on the
common theme. Different species, even in the same
genus, show differences in internal salt concentrations,
degree of osmoregulation, and tolerance of extremes
(e.g., Horne, 1966; Broch, 1988).
The hemoglobins of Moina (a cladoceran), Cyzicus
(Spinicaudata), and Triops (Notostraca) are electrophoretically similar (Horne and Beyenbach, 1974).
Further physiological topics, including diapause,
respiration, and ion balance are discussed in Section
VII. C.
XIV. ECOLOGY OF THE OTHER BRANCHIOPODS
A. Life History
Notostracans begin life as nauplii or metanauplii
which hatch out of eggs, often resting eggs. Non-cladoceran branchiopods use a variety of signals to begin development from the egg, including day length, salinity
level, temperature, and oxygen concentration. Depending on the species, survival through suboptimal conditions is accomplished by either (or both) diapause (an
internal physiological state) or quiescence (slow development resulting from low temperature; Brendonck
1996). There are a dozen or so molts before maturity,
and the adults continue molting throughout their life
(Fryer. 1988).
Anostracans hatch from resting eggs as nauplii or
metanauplii. Some species have the egg shell as nauplii
(Artemia, Anderson, 1967; Branchinecta, Daborn,
1977a), and some hatch as advanced metanauplii with
as many as ten pairs of swimming legs (Eubranchipus,
Daborn, 1976). As the metanauplii molt, they add segments and apendages, gradually developing into adults
(Fryer, 1983). Adults molt several times. Fertilization
normally takes place just after the adult female molts.
The brood pouch has a pore through which sperm are
injected by one or the other of the penes of a male. [It
is not clear how the eggs of Artemiopsis, which lacks a
brood-pouch pore, are fertilized.] Fairy shrimp typically carry eggs until they molt. New eggs are released
into the ovisac only after mating in all genera studied,
except Artemia (Munuswamy and Subramoniam,
1985). Anostracans probably do not store sperm
(Wiman, 1979).
Most conchostracans hatch from resting eggs as
nauplii (Anderson, 1967). The carapace appears in
about the third naupliar stage. By successive molts, the
juveniles gradually come to resemble adults.
894
S. I. Dodson and D. G. Frey
The eggs of most species of large branchiopods enter diapause soon after development commences and
become resistant resting eggs (Belk and Cole, 1975).
Exceptions include Artemia, in which development
may continue in the ovisac and free-swimming nauplii
may pass out the brood pouch, and perhaps Cyclestheria hislopi (Spinicaudata), which may exhibit direct larval development similar to cladocera (D. Belk, personal
communication). Eggs produced parthenogenetically or
sexually have similar diapausing characteristics (Eulimnadia antlei; Belk, 1972). Anostracans, conchostracans,
and notostracans of temporary ponds typically have
only one generation per wet episode and produce resting eggs. There is some evidence that diapausing eggs
of different anotracan species specialize on hatching
under specific rainfall patterns, allowing different
species to take advantage of variations in climatic conditions from year to year in the same temporary pond
(Donald, 1982; Gallagher, 1996).
Thiel (1963) and Belk (1972) raised Triops (Notostraca) and Eulimnadia (Spinicaudata), respectively, in
the laboratory without drying the eggs. The eggs of Triops overwinter best if exposed to low oxygen. The eggs
of Streptocephalus seali show sensitivity to strong drying conditions. This many limit their distribution, excluding them from ephemeral (temporary) ponds in
highly desiccating environments (Belk and Cole, 1975).
Artemia can develop directly in permanent lakes.
Artemia of Mono Lake, a permanent saline lake in the
Great Basin desert of mideastern California, hatch
from resting eggs in the spring, resulting in adults that
produce a second generation from eggs carried and matured in their brood sacs (Lenz, 1984).
The large size achieved by most noncladocerans
requires a longer time to first reproduction, which is
crucial in ephemeral ponds (Loring et al., 1988). Notostracans (1 – 2 cm long) require 2 – 3 weeks to develop from the egg to mature adults (Rzoska, 1961).
Anostracans (1 – 2 cm long) mature in 3 weeks at
15°C (Mossin, 1986) and about 45 days at 4 – 17°C
(Modlin, 1982). Successive generations of Artemia are
separated by a month or so in permanent lakes (Lenz,
1984). Depending on temperature, Eulimnadia diversa (3 – 4 mm) took 4 – 11 days to develop from the
egg to an adult (Belk, 1972). Mature, 1 to 2 mm long
cladocerans, which live in ephemeral ponds (e.g.,
Daphnia, Moina), could produce a clutch of eggs in as
few as 2 days at high temperatures (Rzoska, 1961).
Thus, cladocerans can mature in the most temporary
of ponds, while the larger noncladocerans require
ponds that retain water long enough to allow completion of their longer life cyles. These large branchiopods may then disappear as slower-developing
predators and competitors become abundant (Sublette
and Sublette, 1967; Dodson, 1987; Loring et al.,
1988).
B. Distribution and Biogeography
Because of their vulnerability to fish predation, the
noncladoceran branchiopods typically occur in the absence of fish (Hartland-Rowe, 1972; Kerfoot and
Lynch, 1987). However, in each group, there are a few
reports of occurrences in lakes with fish. Anderson
(1974) gives examples of all three groups in lakes. Some
anostracans co-occur with fish in large, deep freshwater
lakes where fish predation is low (Branchinecta in
Canada, Anderson 1974; Polyartemia in Sweden, Nilsson and Pejler, 1973).
The modern distribution of noncladoceran branchiopods is based on 500 million years of history
(Walossek, 1995). A branchiopodlike animal first appears in the Upper Cambrian. The major groups of
branchiopods date from the Paleozoic, with the exception of cladocerans, which probably diverged from
conchostracans in the Mesozoic or Cretaceous, perhaps
barely 100 million years ago.
Conchostracans are rare north of southern
Canada, while anostracans and notostracans are found
from arctic islands to Central America. All three
groups appear to be especially diverse in the arid southwestern United States. Anostracan diversity in Arizona
depends mainly on two general factors: (1) chemical
heterogeneity among (mostly temporary) habitats; and
(2) thermal variation resulting both from ponds filling
at different seasons and from altitudinal and latitudinal
effects (Belk, 1977). It is uncommon to find congeneric
assemblages within the same pond, although associations of different genera are common (Sublette and
Sublette, 1967; Dodson, 1979, 1987, Donald, 1982;
Kerfoot and Lynch, 1987); however, the two genera of
tadpole shrimp, Triops and Lepidurus, do not appear
to co-occur.
Conservation ecology is an important issue for the
larger branchiopods; hence, a conservation-oriented
newsletter, Anostracan News, is devoted entirely to
them. As its name suggests, the focus is on fairy
shrimp, but it often includes information about large
branchiopods in general. Conservation of large branchiopods is an issue because many of the species have
restricted ranges with only a few small populations in
fragile habitats (such as rock pools in deserts) in danger
of loss or modification. Several large branchiopod
species are listed as endangered species by the U.S.
federal government. As careful studies of species continue (for example, Torrentera and Dodson, 1995) ad-
21. Cladocera and Other Branchiopoda
ditional species with restricted distribution are being
discovered.
C. Physiological Adaptations
Ponds without fish usually exhibit characteristics
that discourage fish; that is, the ponds tend to be small,
isolated, and often have extreme environments that can
be tolerated by branchiopods but not fish. Physiological factors important to the ecology of large branchiopods include high salinity and low oxygen. Because
fishless ponds are typically temporary, large branchiopods also need a strategy, such as diapause, to live
through periods when the pond is dry.
1. Salinity and Temperature
Some of these habitats are saline, and some species
within the anostracan and conchostracan orders show a
high tolerance to salinity (Hartland-Rowe, 1966; 1972;
D’Agostino, 1980). Artemia salina and Branchinecta
campestris are the most tolerant, surviving in waters
several times as saline as seawater. Most fairy shrimp,
conchostracans, and notostracans tolerate a wide range
of salinities; but a few taxa, probably most species of
Eubranchipus and some of Streptocephalus and
Branchinecta, are restricted to low-salinity water. On
the other hand, Artemia are rarely found in water as dilute a seawater (D’Agostino, 1980; Eriksen and Brown,
1980b). The greater salt tolerance of Branchinecta
mackini (Broch, 1988) may provide an occasional
refuge in time or space from its co-occurring congeneric
predator B. gigas.
The distributions of Artemia species and populations within species can be limited by their salinity tolerances (Hartland-Rowe, 1972; Bowen et al., 1985;
Abreu-Grobois, 1987). The viability of several populations in the A. franciscana (Kellogg) superspecies depended on salinity, the concentration of bicarbonate,
and the chloride / sulfate ratio. These differences in
salinity requirements between populations may be a
factor in the process of speciation. Populations in different types of saline lakes are genetically isolated in nature, because nauplii from one source will die in water
from a different type of saline lake (Bowen et al., 1985).
Because temporary ponds can often get quite
warm, these crustaceans must frequently tolerate high
temperatures (Horne, 1971; Hartland-Rowe, 1972;
Eriksen and Brown, 1980a – c). Conchostracans do not
perform well below 10°C, but they can live for at least
a few hours at 30 – 35°C. Fairy shrimp from desert areas tolerate for several hours temperatures as high as
40°C. Several species that tolerate the highest temperatures are eurythermal, in that they can also swim at
895
temperatures near 0°C. Notostracans have thermal tolerances similar to those of fairy shrimp, but there is significant variation among taxa. For example, the higher
temperature tolerance of Branchinecta mackini provides a temporal refuge from its predator Lepidurus
lemmoni (Eriksen and Brown, 1980b).
2. Oxygen
In warm climates, the combination of high temperatures and high productivities can lead to low oxygen
concentrations during the night, or in the deeper levels
of stratified ponds at any time (Horne, 1971; Loring et
al., 1988). Functional hemoglobin has been observed in
some notostracans (Triops), conchostracans (Cyzicus),
and anostracans (Artemia, Horne and Beyenbach,
1974; Streptocephalus mackini, D. Belk, personal communication). All of the larger branchiopods can regulate
their oxygen consumption and live at low oxygen concentrations. In the laboratory, adult conchostracans
(Cyzicus californicus) showed the greatest tolerance to
low oxygen, surviving down to levels of about 0.6 –
0.7 mg / L (roughly 7 – 9% saturation at 20°C) (Eriksen
and Brown, 1980a). Horne (1971) found a natural population of the conchostracan Cyzicus setosa able to tolerate oxygen concentrations of 0.14 mg / L for 2 h with
no significant mortality. Anostracans (Branchinecta
mackini) tolerated oxygen concentrations down to
about 1.4 – 2.8 mg / L (roughly 15 – 30% saturation at
20°C) (Eriksen and Brown, 1980b). Notostracans (Lepidurus lemmoni) have a minimum oxygen tolerance
similar to anostracans, and show the least ability to regulate oxygen consumption at low oxygen concentrations (Eriksen and Brown, 1980c). Oddly enough, a
field study of lethal oxygen thresholds of these animals
revealed that conchostracans (Eulimnadia inflecta) had
a lower tolerance than two anostracan species (Eubranchipus moorei and Streptocephalus seali) (Moore
and Burn, 1968). The apparently greater tolerance of
the anostracans may have been due to the behavioral
tendency of anostracans (but not E. inflecta) to seek the
oxygenated surface of the pond. All three groups show
a maximum rate of weight-specific oxygen consumption
at about 25°C (Eriksen and Brown, 1980a – c).
The saturation level of oxygen concentration is reduced exponentially by salinity. Seawater holds about
20% less oxygen than freshwater at corresponding temperatures and pressures. More saline waters hold even
less oxygen. Broch (1969) suggested that Branchinecta
campestris may be limited to waters of lower salinity
than Artemia salina, both because of salt tolerances and
because Artemia produces hemoglobin, which may be
necessary at high concentrations of salt and the correlated low oxygen concentrations.
896
S. I. Dodson and D. G. Frey
3. Diapause
In Anostraca, Conchostraca, and Notostraca, the
eggs are carried for at least a short period by the adult
female. After a few cell divisions, the egg typically enters diapause. This resting egg (actually a developing
embryo) has a dark covering (tertiary envelope or shell)
and is prepared to survive drying, heat, freezing, and
ingestion by birds (Belk and Cole, 1975). Belk (1970)
removed the covering of diapausing conchostracan embryos to demonstrate that the shell does not reduce
desiccation effects, but may reduce mortality due to
abrasion or intense (ultraviolet) sunlight.
Signals that break diapause include temperature
and oxygen concentration (Hart-Rowe, 1972; Belk and
Cole, 1975; Clegge and Conte, 1980; Wiggins et al.,
1980; Mossin, 1986). In some anostracans, the final signal that breaks diapause may be a combination of high
oxygen and an increase in CO2 concentration (Mossin,
1986). In fact, the fastest average hatching time of the
European anostracan Siphonophanes grubei occurred at
pH of 5.5 (associated with high CO2 concentration). At
least one conchostracan, Caenestheria setosa, has a requirement for high temperatures to break diapause; its
resting eggs hatch between 20 – 35°C. Hatching in Eulimnadia antelei (Spinicaudata) was inhibited by high
salinity and darkness (Belk, 1972).
Resistant eggs appear to be an adaptation to temporary habitats and long-distance dispersal (Belk and
Cole, 1975). The habit of carrying diapausing eggs
probably has two major advantages (Daborn, 1977b).
First, the eggs are protected from numerous egg predators, such as other crustaceans and chironomid larvae.
Second, the eggs are more likely to be ingested by a bird
for long-range dispersal. As with cladocerans and copepods, ingestion of dipausing eggs may be an important
means of dispersal. Proctor (1964) raised Artemia,
Streptocephalus, Triops, and Cyzicus from feces of mallard ducks, Anas platyrhynchos. Yet, curiously, anostracans show subtle genetic differences among populations, even when close to each other, suggesting that the
dispersal rate is actually very low (Boileau et al., 1992).
D. Behavior
1. Swimming Behavior
Notostracans are detritus feeders and predators,
plowing through the superficial sediments to capture
anostracans and benthic invertebrates. When feeding, the
tadpole shrimp is face down and its many legs moving
rhythmically, digging into the sediments, and throwing
up a plume of sediment behind the animal. Most anostracans feed on their backs, filtering with rhythmic movements of their legs as they swim through the water.
Artemiopsis and Branchinecta also roll over and scrape
surfaces with their thoracic legs (Daborn, 1977b).
Branchinecta gigas catches and holds smaller anostracans
with its spiny phyllopods (Fryer, 1966; Daborn, 1975).
Conchostracans swim slowly, spending most of
their time skimming along the pond bottom in aquatic
vegetation (Martin et al., 1986), or above algal mats
(e.g., Sissom, 1980). Notostracans and anostracans will
swim away from a person walking up to the edge of a
pond; conchostracans show no such response.
2. Mating Behavior
Tadpole shrimp are generally considered to be hermaphrodites that need to find a mate for exchange of
sperm. The immediate goal of notostracan mating behavior is for two individuals to join their eleventh pair
of thoracic legs. Male and female gonopores are in the
same position at the base of the first endopodite on
these legs (Akita, 1971), so some bodily contortion is
required to deposit spermatozoa into the brood pouch
of the partner. Engelmann et al. (1996) reported several
populations composed of males and females and suggested that sexual reproduction may be common in notostracans.
Most conchostracan species have even numbers of
males and females, which are morphologically distinct
(Sassaman, 1995). Males have specialized hooks on the
first (Laevicaudata) or first and second (Spinicaudata)
pairs of thoracic legs. These hooks are probably used in
a similar manner to those of cladocerans; the hooks
hold onto the female and position the animals for
ventral apposition copulation (Mathias, 1937; Martin
and Belk, 1988). In laboratory cultures and in nature, it
is common to see a female swimming about with a male
holding on to the lower (posterior) margins of her carapace. In some species, females can fertilize themselves.
Anostracans typically have two sexes, although in
some populations the females reproduce asexually
(Browne, 1993; Badaracco et al., 1995). The sexes are
dimorphic. Males possess enlarged second antennae,
often with long and complex fingerlike appendages.
These antennae are used to grasp the female just anterior to her genital segments (just behind the last thoracic leg). Only the second antennae are prehensile.
Belk (1984) offered evidence indicating no “holding”
role for antennal appendages and suggested none for
frontal appendages. Anostracans may swim in tandem
for days, especially Artemia and Artemiopsis (Daborn,
1977b). Belk (1984) described the mating behavior of
Eubranchipus serratus:
“Once a sexually active male Eubranchipus locates another fairy
shrimp, he positions himself with his head below the dorsal
surface of the other shrimp’s genital segments. From this stationtaking posture (Moore and Ogren, 1962), the male may be able
21. Cladocera and Other Branchiopoda
to assess suitability of mates (Wiman, 1981). Assuming the
station-taking male is below a female, his next action is to clasp
her rapidly just anterior to her genital region and his second
antennae. As he grabs her, he extends his antennal appendages,
laying them along her back. Holding his body stiff with his
phyllopods pressed against the ventral surface of his trunk and
flexing only at the neck region, he bats at the female several
times with quick ventral movements. As soon as she is grabbed,
the female rolls the anterior half of her body into a ball with her
head pressed ventrally against her genital region. Her abdomen
remains straight. This position is usually held only a second or
so. Then the female will either struggle violently and dislodge the
male, or she will relax and begin swimming slowly. If the female
accepts the male, he too relaxes and begins swimming with her
while still clasping her with his second antennae. His antennal
appendages remain stretched along her back. The male now
attempts intromission, trying from first one side, then the other.
Once one of his penes is inserted into the female’s ovisac, the
male assumes an S-posture. He ceases swimming and his
phyllopods make only slight vibration movements. The female
may continue to swim or she may lie on the bottom.
Disengagement is effected by the female with one or more rapid
jerking movements. As soon as copulation terminates, the male
resumes normal swimming.” (p. 69)
Mating in Streptocephalus mackini was similar to
that of E. serratus, except that males took station
above rather than below females (Wiman, 1981). Male
E. serratus were successful in grasping a female in only
about 20% of their attempts (Belk, 1984), whereas S.
mackini males mate with “undiscriminating eagerness,” approaching males as well as females and also
orienting toward fairy shrimp of different genera
(Wiman, 1981). Copulation is terminated by the female, possibly when she releases his penis from her
gonopore (Wiman, 1981). Males are sometimes observed being towed along by their penis and one occasionally finds males with only one functional penis, the
other being everted permanently. The spines on anostracan penes may be used to hold the penis inside the
gonopore during swimming (Wiman, 1981). The
pattern of spines on the penis is a useful taxonomic
character (Brendonck, 1995, 1997).
Eubranchipus bundyi occurs in April in roadside
ditches. Dodson (unpublished) watched four adult
males and one adult female that were swimming in several adjacent depressions in a marshy area near Madison, Wisconsin. Each depression was about 20 cm
deep, with dead grasses at the bottom. One or two
males swam around the edge of each depression, making circles about 20 cm in diameter and keeping at
middepth, or about 10 cm most of the time. Although
both sexes beat their legs 8 – 10 times / s, males swam at
about 1 cm / s, females at 0.1 cm / s. Females swam in
long straight lines, changing direction every minute or
so with a quick burst of speed. They emerged from
grass cover for a few minutes every few hours. A female that came up to the midwater level was followed
897
by a male, who adjusted his swimming rate to hers,
and swam with his head a millimeter or so below her
uropods. He followed for about 10 s, in, then moved
closer. The female escaped with a quick burst of speed,
about 10 cm / s for 4 cm. The male showed no further
interest. The impression these observations give is that
males patrol bits of open water, and females spend
most of their time in concealment. When the females
desire to mate, they swim upward, find a male, mate,
and then return to the protection of the bottom. The
leklike pattern of male and female fairy shrimp mating
behavior may result from their reproductive strategy
(Wiman, 1981). Because females product large clutches
of eggs while males spend little of their energy budget
on gametes, life-history theory predicts that females
should show discrimination in mating while males
should be active and mate as many females as possible.
As a consequence of higher activity levels, males are
more vulnerable than females to predators.
E. Foraging Relationships
In general, anostracans are free-swimming filter
feeders, conchostracans process detritus from surfaces
or collect plankton, and notostracans are benthic detritus feeders tending toward carnivory (Kaestner, 1970).
There are many records of anostracans eating algae.
The filtration mechanisms of the thoracic legs is described by Fryer (1983). Cannon (1933) analyzed the
feeding movements of fairy shrimp legs, and assumed
they lived mainly on algae. However, Bernice (1971) reported that Streptocephalus dichotomus ate a variety of
foods, including large algae (e.g., diatoms and bluegreen bacteria), ciliates, small invertebrates (e.g., rotifers,
nematodes, and cladocerans), and crustacean eggs. The
diet of Artemia is well known both for natural populations and for laboratory, cultures (D’Agostino, 1980).
Artemia feed on coccoid green algae in Mono Lake
(Winkler, 1977), and Baird (1850) relayed an account of
Artemia catching and eating their own nauplii.
Branchinecta gigas, the largest extant anostracan,
catches and eats other smaller fairy shrimp, and possibly
other small zooplankton and benthos (Fryer, 1966;
Daborn, 1975). Several species of Branchinecta and
Artemiopsis (Fryer, 1966; Daborn, 1977b) have the
habit of rolling over and scraping the pond substrate
with their thoracic legs. Several species of Branchinecta
have been seen scooping up detritus and then sorting
through it. Branchinecta species have a row of short
curved spines along the inner distal edge of the legs
which are probably used for scraping. Even congeneric
species show differences in their feeding behavior. Eubranchipus holmni has spinier legs than E. vernalis and
shows more scraping behavior (Modlin, 1982). This
898
S. I. Dodson and D. G. Frey
scraping action seems a preadaption for feeding on benthos and catching larger planktonic prey, methods employed by the predaceous B. gigas.
Conchostracans feed somewhat like cladocerans,
by drawing water into the carapace to remove food
particles using the phyllopods (Kaestner, 1970). They
probably can remove large particles, such as large algal
colonies, small crustaceans, and rotifers. They are able
to feed in soft mud.
Notostracans feed by plowing along the surface or
through muddy sediments. They appear to pump mud
posteriorly and probably extract any organic particles.
Their diet includes algae, amphibian eggs, smaller crustaceans, insect larvae, anostracans, and tadpoles
(Kaestner, 1970; Dodson, 1987). They may be significant predators on resting eggs of other aquatic invertebrates such as fairy shrimp and clam shrimp (Daborn,
1977b). Notostracans capture mosquito larvae, which
is seen as a benefit to humans, but they are also occasionally pests in rice fields, eating the young plants off
at mud level (Dodson, 1987).
Tadpole shrimp, available commercially as dried
eggs, can be easily cultured in the laboratory. Horne
(1966) cultured Triops in the lab using live fairy
shrimp. In the lab, notostracans have been observed to
kill and eat goldfish (C. Sassaman, personal communication).
F. Population Regulation
Cladoceran populations are often controlled by a
combination of physical and biological factors, with
food limitation and predation being important for several generations. Anostracans, conchostracans, and notostracans typically have one generation each time their
habitat appears, in some cases briefly once a year (Belk
and Cole, 1975; Loring et al., 1988). A major strategy
is to make as many small resistant eggs as possible in
the shortest amount of time. These eggs then hatch
when the pond fills again, and the animals typically
show rapid growth due to a combination of high temperatures and physiological capabilities specialized for
rapid growth (Loring et al., 1988). Population regulation can be accomplished by physiologic factors (e.g.,
salinity, Broch, 1988).
In the laboratory when food is not limiting, Artemia
females produced the same total number of eggs with little regard as to whether they are mated early or late in
life (Browne, 1982). They accomplished this by having
larger clutches closer together when mated late in life.
Only at limiting food levels was there a trade-off between adult life span and number of eggs produced.
Populations of Artemia in permanent lakes can be
reduced by bird predation. Mono Lake, a large saline
lake in mideastern California, hosts a dense population
of Artemia. The Artemia occupy the epilimnion (about
15 m deep) from early spring to December. Cooper
et al. (1984) found that the Artemia population declined rapidly in response to feeding by eared grebes
using the lake as a migratory stop in August and September. The Artemia in Mono Lake typically have two
generations per year, with a significant fraction of the
second generation being consumed by grebes. The
Artemia population of Mono Lake lives at great
enough depths to be protected from predation by most
waterfowl, which are surface feeders or shallow divers.
However, waterfowl can be important predators of
large branchiopods in shallow ponds (Dodson and Egger, 1980), and flamingoes are a major predator of
Artemia in shallow tropical ponds and lakes (Hulbert
et al., 1986). Anostracans are an important part of the
diet of many waterfowl, especially dabbling ducks
(Swanson et al., 1985) feeding in midwestern prairie
ponds. Artemia in a permanent lake are a significant
source of food for California gull chicks (Lenz, 1984).
G. Functional Role in the Ecosystem
Because of their large size and abundance, the large
branchiopods living in ephemeral ponds are a major
link between primary production and detritus on the
one hand and predators (often terrestrial) on the other
hand (Loring et al., 1988). Temporary ponds in southwestern North America, especially in physically simple
playas or rock pools, provide community ecologists
with simple, highly replicated, and easily manipulated
natural communities (Dodson, 1987; Loring et al.,
1988). These natural ponds are easily mimicked by
constructing artificial ponds for community studies of
ephemeral pond branchiopods (Tribbey, 1965).
Ephemeral ponds in arid areas are closely linked to
the surrounding terrestrial landscape (Loring et al.,
1988). The ponds are an important water source for
terrestrial animals, which then act as dispersal agents
for crustaceans. Primary production both in the ponds
and on the pond bottoms when they dry are an important food source for terrestrial animals. The position of
Artemia in the food web of a permanent saline lake is
discussed by Winkler (1977).
H. Toxicology
Anostracans, especially species of Artemia and
Streptocephalus, are frequently used in tests of the
acute or chronic effects of environmental contaminants. The Artemia Reference Center of Ghent is a
huge data base, especially for the biology, aquaculture,
and toxicology of Artemia. Endpoints of branchiopod
21. Cladocera and Other Branchiopoda
bioassays are typically survivorship or fecundity (for
example, see Centeno et al., 1993), although more subtle endpoints, such as hatching of cysts (resting eggs) is
also used (Crisinel et al., 1994).
XV. CURRENT AND FUTURE
RESEARCH PROBLEMS
Temporary ponds are a chancy habitat to study,
because they do not always contain water. However, biologists who are willing to work around the vagaries of
these animals and their habitats have an almost unlimited field for genetic, ecological, and behavioral studies
(e.g., Loring et al., 1988). Areas of great potential include application of molecular techniques to questions
of phylogeny and speciation, field studies of patterns of
life-history evolution, grazing and predator – prey relationships, individual behavior, population dynamics,
community structure, and landscape ecology.
XVI. COLLECTING AND REARING TECHNIQUES
Collecting these animals is easy enough when they
are in their adult forms. A large-mesh plankton net, a
D-frame aquatic dip net, or even a large-bore pipette is
sufficient to catch them. The Eubranchipus illustrated
in this chapter were collected with a spoon. A traditional technique is to collect mud from a dried habitat,
add water, and rear the animals that hatch from the re-
899
sistant eggs. Rearing is easy. Anostracans and conchostracans eat commercial tropical fish food. Notostracans
can be fed anostracans, earthworms, or dried cladoceran fish food. Artemia diets are myriad, as are recipes
for culture media. The most common culture medium
is artificial seawater, although more specialized media
may give better viability (Bowen et al., 1985).
XVII. TAXONOMIC KEYS FOR NON-CLADOCERAN
GENERA OF BRANCHIOPODS
A. Available Keys and Specimen Preparation
General keys to North American branchiopod genera can be found in Pennak (1989). More recent revisions of large branchiopod groups include keys for
Streptocephalidae (fairy shrimp, Maeda-Martinez
et al., 1995) and Lynceidae (clam shrimp, Martin and
Belk, 1988).
This key applies to those crustaceans that have
more than ten pairs of thoracic (or trunk) leaflike legs
and paired compound eyes. As in the cladoceran orders, the taxonomy of the larger branchiopods in undergoing revision based on studies of morphology (e.g.,
Brendonck, 1995, 1997, Maeda-Martinez, 1995) and
molecular studies (e.g., Wiman, 1979; Browne and
Sallee, 1984). The animals in this key can be identified
to genus with a good dissecting microscope, without
making dissections. Male or female conchostracans and
notostracans may be identified to genus; males only are
keyed for Anostraca.
B. Taxonomic Key to Genera of Non-Cladoceran Freshwater Branchiopoda
1a.
Body soft and flexible, not covered by a shell (order Anostraca) (for a key to species see Belk, 1975) ................................................3
[Belk and Brtek (1995) prepared a checklist of the global anostracan fauna. They provide a strong argument for conservation of
these animals and their habitat.]
1b.
Body covered at least in part by a hard shell (Figs. 75, and 82) ..........................................................................................................2
2a(1b).
Head and body covered with a clamlike shell that wraps around the legs; the abdomen is enclosed inside the shell (conchostracan
orders Laevicaudata and Spinicaudata) (Figs. 75 and 76) .................................................................................................................13
2b.
Head and body covered dorsally with a flattened shell; the abdomen is straight and trails out behind the shell (order Nostraca)
(Fig. 82) ...........................................................................................................................................................................................20
[Fryer (1988) is an excellent source for notostracan biology and taxonomy.]
3a(1a).
Seventeen pairs of thoracic legs (Fig. 65)........................................................................................................................Polyartemiella
[Belk and Brtek (1995) list a single species for North America.]
3b.
Eleven pairs of thoracic legs ...............................................................................................................................................................4
4a(3b).
Abdomen ends in and is bordered by a flattened blade (Fig. 66A, B) .........................................................................Thamnocephalus
4b.
Abdomen ends in two blade-shaped appendages (cercopods) (Fig. 67) ...............................................................................................5
5a(4b).
Large branched frontal appendage growing from between the second antennae (similar to that of Thamnocephalus, Fig. 66C); when
unrolled, frontal appendage is longer than the second antennae .........................................................................Branchinella (in part)
[Belk and Sissom (1992) report four species of Branchinella from North America. They suggest that at least two of these species
qualify as endangered species, because of habitat destruction.]
900
S. I. Dodson and D. G. Frey
FIGURE 70 Eubranchipus hundyi Forbes, 1876. AB Woods Pond, Dane Co., Wisconsin. 27 April 1986.
(Coll. K. Parejko). (A) Head of male, antennal appendage unrolled; (B) female; (C) male. (KE) FIGURE 71
Dexteria floridanus (Dexter, 1953). Alachua Co., Florida. 7 March 1939. From paratypes in US National Museum. (Collection #93537). (KE) FIGURE 72 Artemiopsis stephanssoni (Johansen, 1922). Head of male.
(Coll. D. Belk #506). (KE) FIGURE 73 Linderiella occidentalis (Dobbs, 1923). Sulfur Mtn. Pond, Venture
Co., California. 2 March 1986. (Coll. S. Copper). (A) Head of male; (B) Male clasping female. (KE) FIGURE
74 Branchinecta coloradensis Packard, 1874. Head of male. Stock Pond in Rock Garden, Grand Co., Utah.
29 October 1986. (KE)
21. Cladocera and Other Branchiopoda
5b.
901
Frontal appendage absent, or if present, unbranched and less than half as long as the basal segment or the second antennae ............6
6a(5b). Lateral spines at the posterior margin of each abdominal segment, spines about 1 / 3 as long as the width of the segment
(Fig. 67B) ............................................................................................................................................................Branchinella (in part)
6b.
Abdomen without lateral spines .........................................................................................................................................................7
7a(6b).
Distal (second) segment of the second antennae a flat triangular blade, the base of the triangle about half of the length (Fig. 68A)
................................................................................................................................................................................................Artemia
[Belk and Brtek (1995) list three species for North America. Hontoria and Amat (1992) used a multivariate analysis of 25 different
North American populations, and found evidence for only two species. However, studies in the Caribbean (Robert Mayer, personal
communication) and Yucatan (Torrentera and Dodson, 1995) suggest the existence of several additional species.]
7b.
Distal segment not triangular and thin, instead more or less rounded, but may be flattened toward the tip (do not confuse the distal
antennal appendage, which is attached on the basal-medial margin of the second antenna and often carried folded or rolled up)
(Fig. 70A) ...........................................................................................................................................................................................8
8a(7b).
Antennal appendages present and longer than the basal (first) segment of the second antennae .........................................................9
8b.
Antennal appendages (projections) on the basal segment, if present, no longer than half of the length of the basal segment of the
second antennae ...............................................................................................................................................................................10
9a(8a).
Complex (branched) medial process attached at the inner corner of the apex of the first segment of the second antennae. The second
segment is also attached at the apex of the basal segment, but it is outside (lateral to) the medial process and unbranched (Fig. 69)
....................................................................................................................................................................................Streptocephalus
[Maeda-Martinez et al. (1995) give a diagnosis and phylogeny of the thirteen North American species of Streptocephalus.]
9b.
Antennal appendage unbranched but with a series of lobes and teeth along at least one margin; antennal appendage attached near
the proximal end of the basal segment of the second antenna ..........................................................................................................11
10a(8b).
Penis with a stout terminal spine about as long as the eversible (distal) part of the penis; several species have a second antenna with
branch-like processes on the second segment (Fig. 70) ....................................................................................................Eubranchipus
[Belk and Brtek (1995) list eight species for North America.]
10b.
Penis without a terminal spine, distal segment of second antenna without branch-like processes (Fig. 71)..............................Dexteria
[According to Belk and Brtek (1995), there is one species in this genus which is closely related to Eubranchipus. The species is
known only from its type locality near Gainesville, FL.]
11a(9b).
Second segment of the second antenna with one or two sharp projections from the medial surface, about halfway along the segment, and approximately doubling the width of the segment at that point; tip of the second segment tapers to a blunt point (Fig. 72)
Artemiopsis
[Belk and Brtek (1995) list a single species for North America, restricted to arctic ponds.]
11b.
Second segment of second antenna with no projections at middle, but sometimes flattened, curved, or with a knob at the tip.........12
12a(11b).
First segment of second antenna with a dorsal-medial projection from the base of the segment about half as long as segment; this is
the only projection on the basal segment (Fig 73) ................................................................................................................Linderiella
[One species is recognized in North America (Belk and Brtek, 1995), and it is a candidate for endangered species status.]
12b.
First segment of the second antenna may be without ornamentation, however, usually there is a projection, bump or patch or row
of spines on the medial surface of the segment from about the middle of the segment, or a ventral-medial projection (curving
upward) from the base of the segment, or some combination of the above; none of the projections more than half as long a the
basal segment (Fig. 74) ....................................................................................................................................................Branchinecta
[Belk and Brtek (1995) list 14 species for northern North America, and two from Mexico. Three of the species found in California
are candidates for listing as endangered species.]
13a(2a).
Dorsal union of the carapace valves a true hinge with a groove (Fig. 75A) (order Laevicaudata) .....................................................19
[Martin and Belk, 1988) provide keys for the family Lynceidae in the Americas.]
13b.
Dorsal union of the valves a simple fold (order Spinicaudata) ..........................................................................................................14
14a(13b).
Dorsal margin of head with a stalked rounded organ (Fig. 76), and an occipital crest at the posterior end of the dorsal margin of the
head .................................................................................................................................................................................................15
14b.
Dorsal margin of head with occipital crest only (Fig. 78), may be indistinct.....................................................................................16
15a(14a).
Ventral surface of telson at point of articulation of terminal claws with a pine (Fig. 76) ....................................................Eulimnadia
[Belk (1989) reviews the North American species of Eulimnadia, recognizing five of the twelve described species as valid, based on
traditional morphologic characters and new information on egg-shell morphology.]
902
S. I. Dodson and D. G. Frey
FIGURE 75
Lycneus brachyurus O.F. Mueller, 1785. Dane Co., Wisconsin. (A) Dorsal groove and hinge; (B)
lateral view; (C) anterior view showing rostrum shape. (KE) FIGURE 76 Eulimnadia cf. agassizzii Packard,
1874. Hwy 190, Oxaca, Mexico. 12 July 1976. (Coll. P. Mündel #235). (KE) FIGURE 77 Limnadia lenticularis (Linnacus, 1761). Female within shell. Woods Hole, MA. 27 August 1887. (USNM coll. Smith). (KE)
FIGURE 78 Leptisthera compleximanus (Packard, 1877). Mexico? (A) Female within shell; (B) Female with
one valve of shell removed. (KE)
21. Cladocera and Other Branchiopoda
FIGURE 79 Eocyzicus concavus (Mackin, 1939). Profile view of male head. Texas. Copied from Mattox,
1950. FIGURE 80 Caenestheriella cf. setosa (Pearse, 1912). (A) Male within shell; (B) male with one valve
removed. (KE) FIGURE 81 Cyzicus californicus (Packard, 1874) Male. Collected east of lordsburg, Hidalgo
Co., New Mexico. 13 August 1955. (Coll Lynch). (KE) FIGURE 82 (A) Lepidurus couessii (Packard, 1875).
Fish hatchery at Valentine, Cherry Co., Nebraska, Spring 1977. (Coll. D.C. Ashley). (KE) (B) Triops longicaudatus LeConte, 1846. Hwy 385, Philips Co., Colorado. 13 June 1972. (Coll. F. Wiman). (KE)
903
904
S. I. Dodson and D. G. Frey
15b.
Ventral surface telson at point of articulation of terminal claws without a spine (Fig. 77).....................................................Limnadia
16a(14b).
Rostrum (snout or beak) with a sharp spine (Fig. 78) ........................................................................................................Leptistheria
16b.
Rostrum with a spine .......................................................................................................................................................................17
17a(16b).
Occipital crest acute, conspicuous ....................................................................................................................................................18
17b.
Occipital crest obtuse, inconspicuous (Fig. 79) .....................................................................................................................Eocyzicus
18a(17a).
Male and female rostrum terminates in an acute point (Fig. 80) ...................................................................................Caenestheriella
18b.
Male rostrum terminates bluntly, anterior margin almost as high as the length of the rostrum (Fig. 81), female rostrum acute
.................................................................................................................................................................................................Cyzicus
19a(13a).
Ridge running from top of head down center of rostrum unbranched; second thoracic leg of male is similar to the more posterior
legs .........................................................................................................................................................................................Lynceus
[Martin et al. (1986) provide comparative descriptions of the North American species, and Martin and Belk (1988) review the family Lynceidae in the Americas.]
19b.
Head ridge branched toward the rostrum; second thoracic leg of male is modified different from the more posterior legs
.........................................................................................................................................................................................Paralimnetis
20a(2b).
Last segment of abdomen (the telson) has a medial extension, the supra-anal plate, between the two long cercopods (Fig. 82A)
............................................................................................................................................................................................Lepidurus
[King and Hannert (1998) present molecular evidence of at least five species of Lepidurus in North America; only four of these have
been named.]
20b.
Telson is concave between the two cercopods (Fig. 82B) ...........................................................................................................Triops
ACKNOWLEDGMENTS
Professor Frey, a meiobenthic specialist, contributed the keys for
the Bosminidae, Chydoridae, and Macrothricidae and discussion of
these families, plus the section on paleolimnology. Professor Dodson
wrote the remaining keys and recently revised the key to the macrothricids and related families. Each author modified the other’s contribution. We are grateful for helpful comments from reviewers, especially
James Thorp, Alan Tessier, Brenda Hann, Denton Belk, Carlos
Santos-Flores, Henri Dumont, Professor Smirnov, and John Havel.
The section on noncladoceran branchiopods could have been written
only with the extensive advice of Denton Belk. Many of the figures for
this chapter were drawn by Kandis Elliot (“KE” in the figure legends).
LITERATURE CITED
Abreu-Grobois, F. A. 1987. A review of the genetics of Artemia, in
Sorgeloos, P., Bengtson, D. A., Decleir, W., Jaspers, E., Eds.,
Artemia research and its applications. Vol. 1. Universa Press,
Wetteren, Belgium, pp. 61 – 73.
Akita, M. 1971. On the reproduction of Triops longicaudatus
(LeConte). Zoological Magazine (Dobutsugaku Zasshi) 80:
242 – 250.
Allen, Y. C., De Stasio, B. T., Ramcharan, C. W. 1993. Individual
and population level consequences of an algal epibiont on
Daphnia. Limnology and Oceanography 38:592 – 601.
Alonso, M. 1996. Crustacea Branchiopoda. Museo Nacional de
Ciencias Naturales, Consejo Superior de Investigaciones Cientificas, Madrid. Fauna Iberica 7:486.
Anderson, D. T. 1967. Larval development and segment formation in
the branchiopod crustaceans Limnadia stanleyana and Artemia
salina (L.) (Anostraca). Australian Journal of Zoology 15:47 – 91.
Anderson, R. S. 1974. Crustacean plankton communities of 340
lakes and ponds in and near the National Parks of the Canadian
Rocky Mountains. Journal of the Fisheries Research Board of
Canada 31:855 – 869.
Badaracco, G., Bellorini, M., Landsberger, N. 1995. Phylogenetic
study of bisexual Artemia using random amplified polymorphic
DNA. Journal of Molecular Evolution 41:150 – 154.
Baird, W. 1850. The natural history of the British Entomostraca.
Printed for the Ray Society, London. reprinted by Johnson
Reprint Corporation, New York. 1968. 364 p 36 tables.
Balcer, M. D., Korda, N. L., Dodson, S. I. 1984. Zooplankton of the
Great Lakes: a guide to the identification and ecology of the
common crustacean species. Uni. of Wisconsin Press, Madison,
WI. 175 pp.
Banta, A. M. 1939. Studies on the physiology, genetics and evolution
of some Cladocera. Paper No. 39, Department of Genetics,
Carnegie Institution of Washington Publication No. 513, pp.
iii – x, 1 – 285.
Bayly, I. A. E. 1986. Aspects of diel vertical migration in zooplankton, and its enigma variations. in: De Deckker P., Williams,
W. D. Eds., Limnology in Australia. CSIRO Melbourne,
Australia, pp. 349 – 368.
Beaton, M. J., P. D. N. Hebert. 1988. Further evidence of hermaphroditism in Lepidurus arcticus (Crustacea, Notostraca) from the
Melville Peninsula area, N. W. T., in: Adams W. P., Johnson, P.
G. Eds., Student Research in Canada’s North: Proceedings of
the National Student’s Conference on Northern Studies. Association of Canadian Universities for Northern Studies, pp
253 – 257.
Belk, D. 1970. Functions of the conchostracan egg shell. Crustaceana
19:105 – 106.
Belk, D. 1972. The biology and ecology of Eulimnadia antlei Mackin
(Conchostraca). The Southwestern Naturalist 16:297 – 305.
Belk, D. 1975. Key to the Anostraca (fairy shrimps) of North America. The Southwestern Naturalist 20:91 – 103.
Belk. D. 1977. Evolution of egg size strategies in fairy shrimps. The
Southwestern Naturalist 22:99 – 105.
Belk, D. 1984. Antennal appendages and reproductive success in the
Anostraca. Journal of Crustacean Biology 4:66 – 71.
21. Cladocera and Other Branchiopoda
Belk, D. 1989. Identification of species in the conchostracan genus
Eulimnadia by egg shell morphology. Journal of Crustacean Biology 9:115 – 125.
Belk, D., Cole, G. A. 1975. Adaptational biology of desert temporary-pond inhabitants. Pages 207 – 226, in: Hadley, N. F., Ed.
Environmental physiology of desert organisms. Dowden,
Hutchinson and Ross, Stroudsburg, Pa.
Belk, D., Brtek, J. 1995. Checklist of the Anostraca. Hydrobiologia
298:315 – 353.
Belk, D., Sissom, S. L. 1992. New Branchinella (Anostraca) from
Texas, USA, and the problem of antenna-like processes. Journal
of Crustacean Biology 12:312 – 316.
Bernardi, R. de. 1984. Methods for the estimation of zooplankton
abundance. Pages 59 – 86, in: Downing, J. A., Rigler, F. H. Eds.
A Manual on methods for the assessment of secondary productivity in fresh waters, 2nd ed. Blackwell Sci., Oxford.
Berner, D. B. 1982. Key to the cladocera of Par Pond on the Savannah River Plant. A publication of the Savannah River Plant, National Environment Research Park Program, U.S. Department
of Energy. NERP-SRO–11. Savanna River Ecology Laboratory,
Aiken, SC.
Bernice, R. 1971. Food, feeding, and digestion in Streptocephalus
dichotomus Baird (Crustacea: Anostraca). Hydrobiologia 38:
507 – 520.
Benzie, J. A. H., Hodges, A. M. A. 1996. Daphnia obtusa Kurz,
1874 ememd. Scourfield, 1942 from Australia. Hydrobiologia
333:195 – 199.
Birge, E. A. 1893. Notes on Cladocera, III. Transactions of the
Wisconsin Academy of Sciences, Arts and Letters 9:275 – 317.
Birge, E. A. 1910. Notes on Cladocera, IV. Transactions of the
Wisconsin Academy of Sciences, Arts and Letters 16:1017 – 1066.
Birge, E. A. 1918. The water fleas (Cladocera), Pages 676 – 750, in:
H. B. Ward, Whipple, G.C. Eds. Fresh-water Biology, Wiley,
New York.
Black, R. W. III, Slobodkin, L. B. 1987. What is cyclomorphosis?
Freshwater Biology (1987) 18:373 – 378.
Boileau, M. G., Hebert, P. D. N., Schwartz, S. S. 1992. Non-equilibrium gene frequency divergency. Persistent founder effects in
natural populations. Journal of Evolutionary Biology 5:25 – 40.
Boucherle, M. M., Zullig, H 1983. Cladoceran remains as evidence
of change in trophic state in three Swill lakes. Hydrobiologia
103:141 – 146.
Bowen, S. T., Buoncristiani, M. R., Carl, J. R. 1988. Artemia habitats: ion concentrations tolerated by one superspecies. Hydrobiologia 158:201 – 214.
Bowen, S. T., Fogarino, E. A., Hitchner, K. N., Dana, G. L., Chow, V.
H. S., Buoncristiani, M. R., Carl, J. R. 1985. Ecological isolation in Artemia: population differences in tolerance of anion
concentrations. Journal of Crustacean Biology 5:106 – 129.
Bowman, T. E. 1971. The case of the nonubiquitous telson and the
fradulent furca. Crustaceana 21:165 – 175.
Bradley, M. C., Perrin, N., Calow, P. 1991. Energy allocation in the
cladoceran Daphnia magna Straus, under starvation and refeeding. Oecologia (Heidelberg) 86:414 – 418.
Brancelj, A. 1990. Alona hercegovinae, new species (Cladocera:
Chydoridae) a blind cave-inhabiting Cladoceran. Hydrobiologia
199:7 – 16.
Brancelj, A. 1992. Alona sketi, new species (Cladocera: Chydoridae)
the second cave-inhabiting cladoceran from former Yugoslavia.
Hydrobiologia 248:105 – 114.
Brandlova, J., Brandl, Z., Fernando, C. H. 1972. The cladocera of
Ontario with remarks on some species and distribution. Canadian Journal of Zoology. 50:1373 – 1403.
Brendonck, L. 1995. An updated diagnosis of the branchipodid genera (Branchiopoda: Anostraca: Branchipodidae) with reflections
905
on the genus concept by Dubois (1988) and the importance of
genital morphology in anostracan taxonomy. Archiv Fuer Hydrobiologie Supplementband 107:149 – 186.
Brendonck, L. 1996. Diapause, quiescence, hatching requirements:
What we can learn from large freshwater branchipods (Crustacea: Branchiopoda: Anostraca, Notostraca, Conchostraca).
Hydrobiologia 320:85 – 97.
Brendonck, L. 1997. The anostracan genus Branchinella (Crustacea:
Branchiopoda), in need of a taxonomic revision: Evidence from
penile morphology. Zoological Journal of the Linnean Society
119:447 – 455.
Bretschko, G. 1969. Zur Ephippienablage bei Chydoridae (Crustacea,
Cladocera). Zool. Anz. Suppl. 33, Vehr. Zool. Ges. 1969:95 – 97.
Brewer, M. C., Coughlin, J. 1996. Virtual plankton: A novel approach to the investigation of aquatic predator-prey interactions. Pages 425 – 434, in: Lenz, P.H., Hartline, D.K., Purcell,
J.E., Macmillan, D.L. editors. Zooplankton: Sensory Ecology
and Physiology. Gordon & Breach Amsterdam.
Brewer, M. C., Dawidowicz, P., Dodson, S.I. 1998. Interactive effects
of fish kairomone and light on behavior in Daphnia. Ecology
Broch, E. S. 1969. The osmotic adaptation of the fairy shrimp
Branchinecta campestris Lynch to saline astatic waters. Limnology and Oceanography 14:485 – 492.
Broch, E. S. 1988. Osmoregulatory patterns of adaptation to inland
astatic waters by two species of fairy shrimps, Branchinecta gigas Lynch and Branchinecta mackini Dexter. Journal of Crustacean Biology 8:383 – 391.
Brooks, J. L. 1957. The systematics of North American Daphnia.
Memoirs of the Connecticut Academy of Arts and Sciences
13:1 – 180.
Brooks, J. L. 1959. Cladocera. Pages 587 – 656, in: W.T. Edmondson,
Editor. Fresh-water Biology, 2nd ed., Wiley, New York.
Browman, H. L., Kruse, S., O’Brien, W. J. 1989. Foraging behavior
of the predaceous cladoceran, Leptodora kindti and escape
responses of their prey. Journal of Plankton Research. 11:
1075 – 1088.
Browne, R. A. 1982. The costs of reproduction in brine shrimp. Ecology 63:43 – 47.
Browne, R. A. 1993. Sex and the single brine shrimp. Natural History 1993 (5):35 – 38.
Browne, R. A., Sallee, S. E. 1984. Partitioning genetic and environmental components of reproduction and lifespan in Artemia.
Ecology 65:949 – 960.
Burns, C. W., Gilbert, J. J. 1986. Effects of daphnid size and density
on interference between Daphnia and Keratella cochlearis.
Limnology and Oceanography 31:848 – 858.
Butorina, L. G. 1986. On the problem of aggregation of planktonic
crustaceans [Polyphemus pediculus (L.), Cladocera]. Archiv fur
Hydrobiologie 105:355 – 386.
Cannon, H. G. 1933. On the feeding mechanism of the branchiopods. Philosophical Transactions of the Royal Society of
London 222:267 – 352
Carpenter, S. (Ed.) 1987. Complex interactions in lake communities.
Springer.
Carpenter, R., Kitchell, J. F., Hodgson, J. R. 1985. Cascading trophic
interactions and lake productivity. BioScience 35:634 – 639.
Carter, J. C. H., Dadswell, M. J., Roff, J. C., Sprules, W. G. 1980.
Distribution and zoogeography of planktonic crustaceans and
dipterans in glaciated eastern North America. Canadian Journal
of Zoology 58:1355 – 1387.
Carvalho, G. R., Hughes, R. N. 1983. The effect of food availability,
female culture-density and photoperiod on ephippia production
in Daphnia magna (Crustacea: Cladocera). Freshwater Biology
13:37 – 46.
Centeno, M. D. F., Brendonck, L., Persoone, G. 1993. Acute toxicity
906
S. I. Dodson and D. G. Frey
tests with Streptocephalus proboscideus (Crustacea: Branchiopoda: Anostraca): influence of selected environmental conditions. Chemosphere 27:2213 – 2224.
Chaplin, J. A., Havel, J. E., Hebert, P. D. N. 1994. Sex and ostracods. Trends in Ecology and Evolution 9:435 – 439.
Cheer, A. Y. L., Koehl, M. A. R. 1987. Paddles and rakes: fluid flow
through bristled appendages of small organisms. Journal of
Theoretical Biology 129:17 – 39.
Chiavelli, D. A., Mills, E. L., Threlkeld, S. T. 1993. Host preference,
seasonality, and community interactions of zooplankton
epibionts. Limnology and Oceanography 38:574 – 583.
Ciros-Perez, J., Elias-Gutierrez. M. 1997. Spinalona anophtalma, n.
gen. n. sp. (Anomopoda, Chydoridae) a blind epigean cladoceran from the Neovolcanic Provence of Mexico. Hydrobiologia 353:19 – 28.
Clegge, J. S., Conte, F.P. 1980. A review of the cellular and developmental biology of Artemia, Pages 11 – 54, in: Persoone, G.,
Sorgeloos, P., Roels, O., Jaspers, E. Eds. The Brine shrimp
Artemia, Vol. 2: Physiology, Biochemistry, Molecular Biology,
Universa Press, Wettern, Belgium.
Colborn, T., Dumanoski, D., Myers, J. P. 1996. Our stolen future.
Dutton. New York. 306 pp.
Colbourne, J. K., Hebert, P. D. N., Taylor, D. J. 1997. Evolutionary
origins of phenotypic diversity in Daphnia. Pages 163 – 188, in:
T. J. Givnish and K. J. Sytsma (Eds.), Molecular evolution and
adaptive radiation. Cambridge Uni. Press.
Cooper, S. D. 1983. Selective predation on cladocerans by common
pond insects. Canadian Journal of Zoology 61:879 – 886.
Cooper, S. D., Winkler, D. W., Lenz, P. H. 1984. The effect of grebe
predation on a brine shrimp population. Journal of Animal
Ecology 53:51 – 64.
Cooper, S. C., Smith, D. W. 1982. Competition, predation and the
relative abundances of two species of Daphnia. Journal of
Plankton Research 4:859 – 879.
Cotten, C. A. 1985. Cladoceran assemblages related to lake conditions in eastern Finland. Ph.D. dissertation, Indiana University,
Bloomington, viii, 96 pp.
Crease, T. J., Hebert, P. D. N. 1983. A test for the production of sexual pheromones by Daphnia magna (Crustacea: Cladocera)
Freshwater Biology 13:491 – 496.
Crisinel, A., Delaunay, L., Rossel, D., Tarradellas, J., Meyer, H.,
Saiah, H., Vogel, P., Delisle, C., Blaise, C. 1994. Cyst-based
ecotoxicological tests using anostracans: Comparison of two
species of Streptocephalus. Environmental Toxicology and
Water Quality 9:317 – 326.
Crisman, T. L. 1980. Chydorid cladoceran assemblages from subtropical Florida. American Society of Limnology and Oceanography Special Symposium 3:657 – 668.
Cummins, K. W., Costa, R. R., Rowe, R. E., Moshiri, G. A., Scanlon,
R. M., Zajdel, R. K. 1969. Ecological energetics of a natural
population of the predaceous zooplankter Leptodora kindtii
Focke (Cladocera). Oikos 20:189 – 223.
Daborn, G. R. 1975. Life history and energy relations of the giant
fairy shrimp Branchinecta gigas Lynch 1937 (Crustacea: Anostraca). Ecology 56:1025 – 1039.
Daborn, G. R. 1976. The life cycle of Eubranchipus bundyi (Forbes)
(Crustacea: Anostraca) in a temporary vernal pond of Alberta.
Canadian Journal of Zoology 54:193 – 201.
Daborn, G. R. 1977a. The life history of Branchinecta mackini
Dexter (Crustacea: Anostraca) in an agrillotrophic lake of
Alberta. Canadian Journal of Zoology 55:161 – 168.
Daborn, G. R. 1977b. On the distribution and biology of an arctic
fairy shrimp Artemiopsis stefanssoni Johansen, 1921 (Crustacea: Anostraca). Canadian Journal of Zoology. 55:280 – 287.
Daday, E. 1898. Mikroskopische Susswasserthiere aus Ceylon.
Terme’s. Fuzetek, Anhangsheft 21:1 – 123.
D’Agusto, A. 1980. The vital requirements of Artemia: physiology
and nutrition. Pages 56 – 82, in: Persoone, G., Sorgeloos, P.,
Roels, O., Jaspers, E. (Eds.) 1980. The brine shrimp Artemia.
Vol. 2. Physiology, Biochemistry, Molecular Biology. Universa
Press: Wetteren, Belgium.
Deevey, E. S. Jr., Deevey, G.B. 1971. The American species of
Eubosmina Seligo (Crustacea, Cladocera). Limnology and
Oceanography 16:201 – 218.
Della Croce, N., Angelino, M. 1987. Marine Cladocera in the Gulf
of Mexico and the Caribbean Sea. Cahiers de Biologie Marine
28:263 – 268.
Della Croce, N., Gaino, E. 1970. Osservazioni sulla biologia del maschio di Penilia avirostris Dana. Cahiers de Biologie Marine
11:361 – 365.
DeMeester, L., Dawidowicz, P., van Gool, E., Loose, C. J. 1998. Ecology and evolution of predator-induced behavior of zooplankton:
Depth selection behavior and diel vertical migration, in: Tollrian,
F., Harvell, C. D., Eds. The ecology and evolution of inducible
defenses. Princeton University Press, Princeton, NJ [in press].
DeMelo, R., Hebert, P. D. N. 1994. Allozyme variation and species
diversity in North American Bosminidae. Canadian Journal of
Fisheries and Aquatic Science 51:873 – 880.
Demott, W. R., Kerfoot, W. C. 1982. Competition among cladocerans: nature of the interaction between Bosmina and Daphnia.
Ecology 63:1949 – 1966.
Dodson, S. I. 1974. Zooplankton competition and predation: An
experimental test of the size-efficiency hypothesis. Ecology
55:605 – 613.
Dodson, S. I. 1975. Predation rates of zooplankton in arctic ponds.
Limnology and Oceanography 20:426 – 433.
Dodson, S. I. 1979. Body size patterns in arctic and temperate zooplankton. Limnology and Oceanography 24:940 – 949.
Dodson, S. I. 1982. Chemical and biological limnology of six westcentral Colorado mountain ponds and their susceptibility to
acid rain. American Midland Naturalist 107:173 – 179.
Dodson, S. I. 1984. Predation of Heterocope septentrionalis on two
species of Daphnia: Morphological defenses and their cost.
Ecology 65:1249 – 1257.
Dodson, S. I. 1985. Daphnia (Ctenodaphnia) brooksi (Crustacea:
Cladocera), a new species from eastern Utah. Hydrobiologia
126:75 – 79.
Dodson, S. I. 1987. Animal assemblages in temporary desert rock
pools: aspects of the ecology of Dasyhelea sublettei (Diptera:
Ceratopogonidae). Journal of the North American Benthological Society 6:65 – 71.
Dodson, S. I. 1989. Predator-induced reaction norms. Bioscience
39:447 – 452.
Dodson, S. 1990. Predicting diel vertical migration of zooplankton.
Limnology and Oceanography 35:1195 – 1200.
Dodson, S., Ramcharan, C. 1991. Size-specific swimming behavior of
Daphnia pulex. Journal of Plankton Research. 13:1367 – 1379.
Dodson, S. I., Egger, D. L. 1980. Selective feeding of red phalaropes
on zooplankton of arctic ponds. Ecology 61:755 – 763.
Dodson, S. I., Dodson, V. E. 1971. The diet of Ambystoma tigrinum
larvae from Western Colorado. Copeia 1971:614 – 624.
Dodson, S. I., Merritt, C. M., Shannahan, J.-P., Shults, C. M. 1999.
Low doses of Atrazine increase male production in Daphnia pulicaria. Environmental Toxicology and Chemistry 18: [in press].
Dodson, S. I, Tollrian, R., Lampert, W. 1997a. Daphnia swimming
behavior during vertical migration. Journal of Plankton Research. 19:969 – 978.
Dodson, S. I., Arnot, S. E., Cottingham, K. L. 1999. The relationship
21. Cladocera and Other Branchiopoda
in lake communities between primary productivity and species
richness. Ecology [in review]
Dodson, S. I., Ryan, S., Tollrian, R., Lampert, W. 1997b. Individual
swimming behavior of Daphnia: effects of food, light, and container size on four clones. Journal of Plankton Research.
19:1537 – 1552.
Dodson, S. I., Hanazato, T., Gorski, P. R. 1995. Behavioral responses
of Daphnia pulex exposed to carbaryl and Chaoborus kariomone. Environmental Toxicology and Chemistry 14:43 – 50.
Dodson, S. I., Hanazato, T. 1995. Commentary on effects of anthropogenic and natural organic chemicals on development, swimming behavior, and reproduction of Daphnia, a key member of
aquatic ecosystems. Environmental Health Perspectives 103
(Suppl. 4):7 – 11.
Donald, D. B. 1982. Erratic occurrence of anostracans in a temporary
pond: colonization and extinction or adaptation to variations in
annual weather. Canadian Journal of Zoology 61:1492 – 1498.
Doolittle, A. A. 1911. Descriptions of recently discovered cladocera
from New England. Proceedings of the U.S. National Museums.
41:161 – 170.
Downing, J. A. 1984. Sampling the benthos of standing waters. Pages
87 – 130, in: Downing, J. A., Rigler, F. H., Eds. A Manual on
Methods for the Assessment of Secondary Productivity in Fresh
Waters, 2nd ed. Blackwell Sci., Oxford. 501 pp.
Dumont, H. J. 1987. Groundwater Cladocera: A synopsis. Hydrobiologia 145:169 – 173.
Dumont, H. J., Pensaert, J. 1983. A revision of the Scapholeberinae
(Crustacea: Cladocera). Hydrobiologia 100:3 – 45.
Dumont, H. J. 1995. The evolution of groundwater Cladocera. Hydrobiologia 307:69 – 74.
Dumont, H.J., Silva-Briano, M. 1998. A reclassification of the
anomopod families Macrothricidae and Chydoridae, with the
creation of a new suborder, the Radopoda (Crustacea: Branchiopoda). Hydrobiologia 384:119 – 149.
Edmondson, W. T. 1960. Reproductive rates of rotifers in natural
populations. Memorie dell’Istituto Italiano di Idrobiologia
12:21 – 77.
Edmondson, W. T. 1963. Pacific Coast and Great Basin. Pages
371 – 392 in: D. G. Frey, Ed. Limnology in North America,.
Univ. of Wisconsin Press. Madison, WI.
Edmondson, W. T. 1991. The uses of ecology: Lake Washington and
beyond. Uni. of Washington Press, Seattle, WA.
Edmondson, W. T., Litt, A. H. 1982. Daphnia in Lake Washington.
Limnology and Oceanography 27:272 – 293.
Edmondson, W. T., Litt, A. H. 1987. Conochilus in Lake Washington. Hydrobiologia 147:157 – 162.
Engelmann, M. Hoheisel, G., Hahn, T., Joost, W., Vieweg, J.,
Naumann, W. 1996. Populations of Triops cancriformis (Bosc)
(Notostraca) in Germany north of 50 degrees N are not clonal
and at best facultatively hermaphroditic. Crustaceana (Leiden)
69:755 – 768.
Engle, D. L. 1985. The production of hemoglobin by small pond
Daphnia pulex: Intraspecific variation and its relation to habitat. Freshwater Biology 15:631 – 638.
Enserink, E. L., Van Der Hoeven, N., Smith, M., Van Der Klis,
C. M., Van Der Gaag, M. A. 1996. Competition between cohorts of juvenile Daphnia magna: A new experimental model.
Archiv für Hydrobiologie 136:433 – 454.
Eriksen, C. H., Brown, R. J. 1980a. Comparative respiratory physiology and ecology of phyllopod crustacea. I. Conchostraca.
Crustaceana 39:1 – 10.
Ericksen, C. H., Brown, R. J. 1980b. Comparative respiratory physiology and ecology of phyllopod crustacea. II. Anostraca. Crustaceana 39:11 – 21.
907
Eriksen, C. H., Brown, R. J. 1980c. Comparative respiratory physiology and ecology of phyllopod crustacea. III. Notostraca. Crustaceana 39:22 – 32.
Ferrari, D. C., Hebert, P. D. N. 1982. The induction of sexual reproduction in Daphnia magna: genetic differences between arctic
and temperate populations. Canadian Journal of Zoology
60:2143 – 2148.
Forbes, S. A. 1925. The lake as a microcosm. Bulletin of the Illinois
State Laboratory of Natural History (Survey) 15:537 – 550.
Fretwell, S. J. 1987. Food chain dynamics: the central theory of ecology? Oikos 50:291 – 301.
Frey, D. G. 1962. Cladocera from the Eemian Interglacial of Denmark. Journal of Paleontology 36:1133 – 1154.
Frey, D. G. 1964. Remains of animals in Quaternary lake and bog
sediments and their interpretation. Archiv für Hydroliologie,
Supplement Ergebnisse der Limnologie 2:1 – 116.
Frey, D. G. 1965. Differentiation of Alona costata Sars from two related species (Cladocera, Chydoridae). Crustaceana 8:159 – 173.
Frey, D.G. 1966. Phylogenetic relationships in the family Chydoridae. Marine Biology Association India, Proceedings of a Symposium on Crustacea, Part 1:29 – 37.
Frey, D. G. 1969. Symposium on paleolimnology. International Association of Theoretical and Applied Limnology, Mitteilungen.
17:1 – 448.
Frey, D. G. 1971. Worldwide distribution and ecology of Eurycercus and
Saycia (Cladocera). Limnology and Oceanoraphy. 16: 254–308.
Frey, D. G. 1974. Paleolimnology. 95 – 123 in Jubilee Symposium: 50
years of limnological research. W. Rodhe (Ed.). International
Association of Theoretical and Applied Limnology, Mitteilungen. 20:1 – 402.
Frey, D. G. 1975. Subgeneric differentiation within Eurycercus
(Cladocera, Chydoridae) and a new species from northern Sweden. Hydrobiologia 46:263 – 300.
Frey, D. G. 1976. Interpretation of Quaternary paleoecology from
Cladocera and midges, and prognosis regarding usability of
other organisms. Canadian Journal of Zoology 54:2208 – 2226.
Frey, D. G. 1978. A new species of Eurycercus (Cladocera, Chydoridae) from the southern United States. Tulane Studies Zoology
& Botany 20:1 – 26.
Frey, D. G. 1980a. On the plurality of Chydorus sphaericus (O. F.
Muller)(Cladocera, Chydoridae) and designation of a neotype
from Sjaelso, Denmark. Hydrobiologia 69:83 – 123.
Frey, D. G. 1980b. The non-swimming chydorid Cladocera of wet
forests, with descriptions of a new genus and two new species.
Internationale Revue der gesamten. Hydrobiologie 65:613 – 641.
Frey, D. G. 1982. Cladocera. Pages177 – 185, in: Hulbert, S. H.,
Villalobos-Figueroa, A. Eds. Aquatic Biota of Mexico, Central
America and the West Indies. San Diego State University, San
Diego, CA.
Frey, D. G. 1982a. Questions concerning cosmopolitanism in Cladocera. Archiv für. Hydrobiol. 93:484 – 502.
Frey, D. G. 1982b. Relocation of Chydorus barroisi and related
species (Cladocera, Chydoridae) to a new genus and descriptions of two new species. Hydrobiologia 86:231 – 269.
Frey, D. G. 1982c. The reticulated species of Chydorus (Cladocera,
Chydoridae): two new species with suggestions of convergence.
Hydrobiologia 93:255 – 279.
Frey, D. G. 1982d, Cladocera. Pages 177 – 185, in: Hulbert, S. H.,
Villalogos-Fiqueroa, A. Eds. Aquatic Bioat of Mexico, Central
America, and the West Indies. SanDiego State Uni. Press, San
Diego, CA.
Frey, D. G. 1985. Cladocera. Proceedings of the Cladocera Symposium,
Budapest 1985, Forro’, L., Frey, D. G., Eds. Junk: Dordrecht.
Frey, D. G. 1986a. The non-cosmopolitanism of chydorid Cladocera:
implications for biogeography and evolution, Pages 237 – 256,
908
S. I. Dodson and D. G. Frey
in: Gore, R. H., Heck, K. L., Eds. Crustacean Biogeography.
Balkema: Rotterdam.
Frey, D. G. 1986b. Cladocera analysis, in: Berglund, B. E. Ed. Handbook of Holocene Palaeoecology and Palaeohydrology. Wiley,
Chicester. pp. 667 – 692,
Frey, D. G. 1987a. The taxonomy and biogeography of the Cladocera. Hydrobiologia 145:5 – 17.
Frey, D. G. 1987b. The North American Chydorus faviformis
(Cladocera, Chydoridae) and the honeycombed taxa of other
continents. Philosophical Transactions of the Royal Society of
London B315:353 – 402.
Frey, D. G. 1988a. Cladocera. Chapter, in: Higgins, R.P., Thiel,
Hjalmar. Eds. Introduction to the study of meiofauna. Smithsonian Press. Washington, DC.
Frey, D. G. 1988b. Are there tropicopolitan macrothricid Cladocera?
Acta Limnologica Brasiliensia 2:513 – 525.
Frey, D. G. 1988c. Separation of Pleuroxus laevis Sars, 1961, from
two resembling species in North America: Pleuroxus straminius
Birge, 1879 and P. chiangi n. sp. (Cladocera, Chydoridae).
Canadian Journal of Zoology 66:2534 – 2563.
Frey, D. G. 1989. Separation of Pleuroxus laevis Sars, 1961, from
two resembling species in North America — P. straminius Birge,
1879, and P. chiangi n. sp. (Cladocera, Chydoridae). Canadian
Journal of Zoology. 66:2534 – 2563.
Frey, D. G., Hann, B. J. 1985. Growth in cladocera. Pages 315 – 335,
in: A. W. Wenner, Ed. Factors in adult growth. Balkema, Boston.
Fryer, G. 1966. Branchinecta gigas Lynch, a non-filter-feeding raptatory anostracan, with notes on the feeding habits of certain
other anostracans. Proceedings of the Linnean Society of
London 177:19 – 34.
Fryer, G. 1968. Evolution and adaptive radiation in the Chydoridae
(Crustacea: Cladocera): a study in comparative functional morphology and ecology. Philosophical Transactions of the Royal
Society of London, Series B, Biological Sciences 254:221 – 385.
Fryer, G. 1970. Defecation in some macrothricid and chydorid cladocerans. Zoological Journal of the Linnean Society 49:255 – 269.
Fryer, G. 1971. Allocation of Alonella acutirostris (Birge) (Cladocera,
Chydoridae) to the genus Disparalona. Crustaceana 21:
221 – 222.
Fryer, G. 1972. Observations on the ephippia of certain macrothricid
cladocerans. Zoological Journal of the Linnean Society 51:
79 – 96.
Fryer, G. 1974. Evolution and adaptive radiation in the Macrothricidae (Crustacea: Cladocera): a study in comparative functional
morphology and ecology. Philosophical Transactions of the Royal
Society of London, Series B, Biological Sciences 269:137 – 274.
Fryer, G. 1983. Functional ontogenetic changes in Branchinecta ferox
(Milne-Edwards) (Crustacea:Anostraca). Philosophical Transactions of the Royal Society of London, Series B. 303:229 – 243.
Fryer, G. 1985. Crustacean diversity in relation to the size of water
bodies: some facts and problems. Freshwater Biology 15:
347 – 361.
Fryer, G. 1987. A new classification of the branchiopod Crustacea.
Zoological Journal of the Linnean Society 91:357 – 383.
Fryer, G. 1988. Studies on the functional morphology and biology of
the Notostraca. Philosophical Transactions of the Royal Society
of London. B. 321(1203):27 – 124.
Fryer, G. 1991. Functional morphology and the adaptive radiation of
the Daphniidae (Branchiopoda: Anomopoda). Philosophical
Transactions of the Royal Society of London. Series B.
331:1 – 99.
Gabriel, W., Taylor, B. E., Kirsch-Prokosch, S. 1987. Cladoceran
birth and death rates estimates: experimental comparisons of
egg-ratio methods. Freshwater Biology 18:361 – 372.
Galat, D. L., Robinson, R. 1983. Predicted effects on increasing
salinity on the crustacean zooplankton community of Pyramid
Lake, Nevada. Hydrobiologia 105:115 – 131.
Gallagher, S. P. 1996. Seasonal occurrence and habitat characteristics
of some vernal pool branchiopoda in northern California,
U.S.A. Journal of Crustacean Biology 16:323 – 329.
Gerritsen, J., Porter, K. G. 1982. The role of surface-chemistry in filter feeding by zooplankton. Science 216:1225 – 1227.
Gerritsen, J., Porter, K. G., Strickler, J. R. 1988. Not by sieving alone:
Suspension feeding in Daphnia. Bulletin of Marine Science
43:366 – 376.
Gorski, P. R., Dodson, S. I. 1996. Free-swimming Daphnia can avoid
following Stokes’ law. Limnology and Oceanography 41:
1815 – 1821.
Gould, S. J. 1989. Wonderful life. Norton, NY.
Gophen, M., Geller, W. 1984. Filter mesh size and food particle uptake by Daphnia. Oecologia (Berlin) 64:408 – 412.
Goulden, C. E. 1966. The animal microfossils. Pages 84 – 120 in:
Cogill, U. M., Goulden, C. E., Hutchinson, G. E., Patrick, R.,
Racek, A. A., Tsukada, M. (Eds), The history of Laguna de
Petenxil. Memoirs of the Connecticut Academy of Arts and
Sciences Vol. 17.
Goulden, C. E. 1968. The systematics and evolution of the Moinidae.
Transactions of the American Philosophical Society, Vol. 58,
New Series, Part 6. pp. 1 – 101.
Goulden, C. E. 1969. Developmental phases of the biocenosis. Proceedings of the National Academy of Sciences 62:1066 – 1073.
Graham, D. M., Sprules, W. G. 1992. Size and species selection of
zooplankton by larval and juvenile walleye (Stizostedion vitreum vitreum) in Oneida Lake, New York. Canadian Journal of
Zoology 7-:2059 – 2067.
Hairston, N. G., Jr. 1996. Zooplankton egg banks as biotic reservoirs in changing environments. Limnology and Oceanography
41:1087 – 1092.
Hall, D. J. 1964. An experimental approach to the dynamics of a
natural population of Daphnia galeata mendotae. Ecology
45:94 – 112.
Hanazato, T., Dodson, S. I. 1993. Morphological responses of four
species of cyclomorphic Daphnia to a short-term exposure to
the insecticide carbaryl. Journal of Plankton Research 15:
1087 – 1095.
Hann, B. J. 1982. Two new species of Eurycercus (Bullatifrons) from
Eastern North America (Chydoridae, Cladocera). Taxonomy, ontogeny, and biology. Int. Rev. ges. Hydrobiology. 67:585 – 610.
Hann, B. J. 1984. Influence of temperature on life-history characteristics of two sibling species of Eurycercus (Cladocera, Chydoridae) Canadian Journal of Zoology 63:891 – 898.
Hann, B. J. 1985. Influence of temperature on life-history characteristics of two sibling species of Eurycercus (Cladocera, Chydoridae). Canadian Journal of Zoology 63:891 – 898.
Hann, B. J. 1986. Revision of the genus Daphniopsis Sars, 1903
(Cladocera: Daphniidae) and a description of Daphniopsis
chilensis, new species, from South America. Journal of Crustacean Biology 6:246 – 263.
Hann, B. J., Chengalath, R. 1981. Redescription of Alonella pulchella Herrick, 1884 (Cladocera, Chydoridae), and a description of the male. Crustaceana 41:249 – 262.
Hann, B. J., Hebert, P. D. N. 1986. Genetic variation and population
differentiation in species of Simocephalus (Cladocera, Daphniidae). Canadian Journal of Zoology 64:2246 – 2256.
Hartland-Rowe, R. 1966. The fauna and ecology of temporary pools
in Western Canada. Verhandlungen. Internationale Vereinigung
für Theoretische und Angewandte Limnologie 16:577 – 584.
Hartland-Rowe, R. 1972. The limnology of temporary waters and
21. Cladocera and Other Branchiopoda
the ecology of Euphyllopoda. Pages 15 – 30, in: Clark, R.B.,
Wootton, R.J., Eds. Essays in Hydrobiology presented to Leslie
Harvey. University of Exeter.
Havel, J. E. 1985. Predaton of common invertebrate predators on
long- and short-featured Daphnia retrocurva. Hydrobiologia
124:141 – 149.
Havel, J. E., Hebert, P. D. N. 1989. Apomictic parthenogenesis and
genotypic diversity in Cypridopsis vidua (Ostracoda: Cyprididae). Heredity 62:383 – 392.
Havel, J. E., Mabee, W. R., Jones, J. R. 1995. Invasion of the exotic
cladoceran Daphnia lumholtzi into North American reservoirs.
Canadian Journal of Fisheries and Aquatic Sciences 52:151 – 160.
Hebert, P. D. N. 1978. The population biology of Daphnia (Crustacea, Daphnidae). Biological Review 53:387 – 426.
Hebert, P. D. N. 1985. Interspecific hybridization between cyclic
parthenogens. Evolution 39:216 – 220.
Hebert, P. D. N. 1987. Genotypic characteristics of the Cladocera.
Hydrobiologia 145:183 – 193.
Hebert, P. D. N. 1988. The comparative evidence. Pages 175 – 196,
in: S. C. Stearns, ed. The evolution of sex and its consequences.
Birkhauser, Boston.
Hebert, P. D. N. 1995. The Daphnia of North America — An illustrated fauna. Version 1. CD ROM.
Hebert, P. D. N., Finston, T. L. 1996. A taxonomic reevaluation of
North American Daphnia (Crustacea: Cladocera). II. New
species in the Daphnia pulex group from the south-central United
States and Mexico. Canadian Journal of Zoology 74:632 – 653.
Hirvonen, H., Ranta, E. 1996. Prey to predator size ratio influences
foraging efficiency of larval Aeshna juncea dragonflies. Oecologia (Berlin) 106:407 – 415.
Hontoria, F., Amat, F. Morphological characterization of adult
Artemia (Crustacea, Branchiopoda) from different geographical
origins: American populations. Journal of Plankton Research
14: 1461 – 1471.
Horne, F. R. 1966. Some aspects of ionic regulation in the tadpole
shrimp Triops longicaudatus. Comparative Biochemistry and
Physiology 19:313 – 316.
Horne, F. R. 1971. Some effects of temperature and oxygen concentration on phyllopod ecology. Ecology 52:343 – 347.
Horne, F. R., Beyenbach, K. W. 1974. Physio-chemical features of hemoglobin of the Crustacean Triops longicaudatus. Archives of
Biochemistry and Biophysics 161:369 – 374.
Hobaek, A., Larsson, P. 1990. Sex determination in Daphnia magna.
Ecology 71:2255 – 2268.
Hrbácek, J. 1958. Density of the fish population as a factor influencing the distribution and speciation of the species of Daphnia.
15th Proceedings of the International Congress of Zoology, Section 10:794 – 796.
Hurlbert, S. H., Loayza, W., Moreno. T. 1986. Fish – flamingo – plankton interactions in the Peruvian Andes. Limnology and Oceanography 31:457 – 468.
Hurtubise, R. D., Havel, J. E. The effects of ultraviolet-B radiation
on freshwater invertebrates: Experiments with a solar simulator.
Limnology and Oceanography 43:1082 – 1088.
Hutchinson, G. E. 1967. A treatise on limnology, Vol. II, Introduction to lake biology and the limnoplankton. Wiley, New York.
1115 pp.
Hutchinson, G. E., Bonatti, E. Cogill, U. M., Goulden, C. E., Leventhall, E. A., Mallett, M. E. Margaritora, F., Patrick, R., Racek,
A., Roback, S. A., Stella, E., Ward-Perkins, J. B., Wellman, T.
1970. Ianula: An account of the history and development of the
Lago di Monterosi, Latium, Italy. Transactions of the American
Philosophical Society. 60 (New Series): pt. 4:3 – 178.
Hyman, L. H. 1926. Note on the destruction of Hydra by a chydorid
909
cladoceran, Anchistropus minor Birge. Transactions of the
American Microscopical Society 45:298 – 301.
Innes, D. J., Hebert, P. D. N. 1988. The origin and genetic basis of
obligate parthenogenesis in Daphnia pulex. Evolution 42:
1024 – 1035.
Jack, J. D., Thorp, J. H. 1995. Daphnia lumholtzi: appearance and
likely impacts of an exotic cladoceran in the Ohio River. Transactions of the Kentucky Academy of Science. 56(3 / 4):101 – 103.
Jurine, L. 1820. Histoire des Monocles, qui se trouvent aux environs
de Geneve. J. J. Paschoud, Paris, 260 pp., 22 plates.
Jensen, K. H., Jakobsen, P. J., Kleiven, O. T. 1998. Fish kairomone
regulation of internal swarm structure in Daphnia pulex
(Cladocera: Crustacea). Hydrobiologia [In Review].
Karabin, A. 1974. Studies on the predatory role of the cladoceran,
Leptodora kindtii (Focke), in secondary production of two
lakes with different trophy. Ekologia Polska 22:295 – 310.
Kaestner, A. 1970. Invertebrate Zoology. Vol. 3. Crustacea [English
edition] Wiley Interscience, New York. 523 pp.
Kerfoot, W. C. 1981. Long-term replacement cycles in cladoceran
communities: a history of predation. Ecology 62:216 – 233.
Kerfoot, W. C., Lynch, M. 1987. Branchiopod Communities: Associations with planktivorous fish in time and space. Pages
367 – 378, in: Kerfoot, W. C., Sih, A., Eds. Predation: Direct and
indirect impacts on aquatic communities. Uni. Press of New
England. Hanover.
Kerfoot, W. K., Sih, A. (Eds.) 1987. Predation: direct and indirect impacts on aquatic communities. Uni. Press of New England.
Hanover. 386 pp.
Kerfoot, W. C., Kellogg, D. L., Jr. Strickler, J. R. 1980. Visual observations of live zooplankters: Evasion, escape, and chemical
defenses. American Society of Limnology and Oceanography
Special Symposium 3:10 – 27.
Kilham S. S., Kreeger D. A., Lynn S. G., Goulden C. E., Herrera L.
1998. COMBO: A defined freshwater culture medium for algae
and zooplankton. Hydrobiologia 377:147 – 159.
King, C. R., Greenwood, J. G. 1992. The productivity and carbon
budget of a natural population of Daphnia lumholtzi Sars.
Hydrobiologia 231:197 – 207.
King, J. L., Hannert, F. 1998. Cryptic species in a “living fossil” lineage: Taxonomic and phylogenetic relationships within the
genus Lepidurus (Crustacea: Notostraca) in North America.
Molecular Phylogenetics and Evolution 10:23 – 36.
Kitchell, J. F. 1992. Food Web Management: A case study of Lake
Mendota. Springer, New York. 553 pp.
Kitchell, J. F., Carpenter, S. A. 1987. Piscivores, planktivores, fossils,
and phorbins. Pages 132 – 146, in: W. C. Kerfoot, Ed. Predation:
Direct and indirect impacts on aquatic communities. New England Press, Hanover, N.H. 386 pp.
Kleiven, O. T., Larsson, P., Hobaek, A. 1992. Sexual reproduction in
Daphnia magna requires three stimuli. Oikos 65:197 – 206.
Klekowski, R. Z. (Ed.) 1978. (Proceedings) Polskie Archiwum Hydrobiologii 25:1 – 501.
Kokkinn, M. J., Williams, W. D. 1987. Is ephippial morphology a
useful taxonomic descriptor in the Cladocera? An examination
based on a study of Daphniopsis (Daphniidae) from Australian
salt lakes. Hydrobiologia 145:67 – 73.
Kořínek, B. 1981. Diaphanosoma birgei n.sp. (Crustacea,
Cladocera). A new species from America and its widely
distributed subspecies Diaphanosoma birgei ssp. lacustris
n. ssp. Canadian Journal of Zoology 59: 1115 – 1121.
Korovchinsky, N. M. 1992. Sididae and Holopediidae. Guides to the
identification of the macroinvertebrates of the continental
waters of the world, No. 3. SPB Academic Publishing. The
Hague. 77 p.
910
S. I. Dodson and D. G. Frey
Kotov, A. A., Boikova, O. S. 1998. Comparative analysis of the late
embryogenesis of Sida crystallina (O.F. Muller, 1776) and Diaphanosoma brachyurum (Lievin, 1848) (Crustacea: Branchiopoda: Ctenopoda) Hydrobiologia [In Press].
Kubersky, E. S. 1977. Worldwide distribution and ecology of Alonopsis (Cladocera: Chydoridae) with a description of Alonopsis
americana sp. nov. Int. Rev. ges. Hydrobiol. 62:649 – 685.
Krueger, D. A., Dodson, S. I. 1981. Embryological induction and predation ecology in Daphnia pulex. Limnology and Oceanography 26:219 – 223.
Lampert, W. 1984. The measurement of respiration. Pages 413 – 468,
in: Downing, J. A., Rigler, F. H. Eds. 1984. A manual on methods for the assessment of secondary productivity in fresh waters, 2nd edn. Blackwell Sci., Oxford.
Lampert, W. 1986a. Response of the respiratory rate of Daphnia
magna to changing food conditions. Oecologia (Berlin)
70:495 – 501.
Lampert, W. 1986b. Phytoplankton control of grazing zooplankton:
A study on the spring clear-water phase. Limnology and
Oceanography 31:478 – 490.
Lampert, W. 1987. Vertical migration of freshwater zooplankton: Indirect effects of vertebrate predators on algal communities.
Pages 291 – 299, in: Kerfoot, W.C. Ed. Predation: direct and indirect impacts on aquatic communities. New England Press,
Hanover, NH.
Lampert, W. 1993. Ultimate causes of diel vertical migration of zooplankton: new evidence for the predator-avoidance hypothesis.
Archiv für Hydrobiologie. Beih. Ergebn. Limnol. 39:79 – 88.
Lampert, W., Sommer, U. 1997. Limnoecology: the ecology of lakes
and streams, (Translated by J. Haney) Oxford Uni. Press.
Landon, M. S., Stasiak, R. H. 1983. Daphnia hemoglobin concentration as a function of depth and oxygen availability in Arco
Lake, Minnesota. Limnology and Oceanography 28:731 – 737.
Larsson P., Dodson, S. I. 1993. Invited review. Chemical communication in planktonic animals. Archiv für Hydrobiologie 129:
129 – 155.
Lehman, J. T. 1987. Palearctic predator invades North American
Great Lakes. Oecologia (Berlin) 74:478 – 480.
Lehman, N., Pfrender, M. E., Morin, P. A. Crease, T. J, Lynch, M.
1995. A hierarchical molecular phylogeny within the genus
Daphnia. Molecular Phylogenetics and Evolution 4:395 – 407.
Lenz, P. H. 1984. Life-history analysis of an Artemia population in a
changing environment. Journal of Plankton Research 6:967 – 983.
Lilljeborg, W. 1901. Cladocera Sueciae, oder Beitraage zur Kenntniss
der in Schweden lebenden Krebsthiere von der Ordnung der
Branchiopoden und der Unterordnung der Cladoceren. Nova
Acta reg. Soc. Sci. Upps. Ser. 3, pp. i-vi, 1 – 701.
Linder, F. 1941. Contributions to the morphology and the taxonomy
of the Branchiopoda Anostraca. Zoologiska Bidrag fran Uppsala 10:101 – 302, 1 plate.
Löffler, H., Ed. 1987. Paleolimnology IV. Proceedings of the Fourth
International Symposium on Paleolimnology, held at Ossiach,
Carinthia, Austria. Hydrobiologia Vol. 143, 431 pp.
Longhurst, A. R. 1954. Reproduction in Notostraca (Crustacea).
Nature 173:781 – 782.
Loose, C. 1993. Lack of endogenous rhythmicity in Daphnia diel vertical migration. Limnology and Oceanography 38:1837 – 1841.
Loring, S. J., MacKay, W. P., Whitford, W. G. 1988. Ecology of small
desert playas. Pages 89 – 113, in: Thame, J.L., Ziebel, C. D. Eds.
Small water impoundments in semi-arid regions. Uni. of New
Mexico Press, Albuquerque.
Luecke, C., Litt, A. H. 1987. Effects of predation by Chaoborus
flavicans of Lake Lenore, Washington. Freshwater Biology 18:
185 – 192.
Luecke, C., O’Brien, W. J. 1983. Photoprotective pigments in a pond
morph of Daphnia middendorffiana. Arctic 36:365 – 368.
Lynch, M. 1980. The evolution of cladoceran life histories. The
Quarterly Review of Biology 55:23 – 42.
Lynch, M. 1982. How well does the Edmondson – Paloheimo model
approximate instantaneous birth rates? Ecology 63:12 – 18.
Lynch, M. 1983. Ecological genetics of Daphnia pulex. Evolution 37:
358 – 374.
Lynch, M. 1989. The life history consequences of resource depression
in Daphnia pulex. Ecology 70: 246 – 256.
MacIsaac, H. J., Hebert, P. D. N., Schwartz, S.S. 1985. Inter- and intraspecific variation in acute thermal tolerance of Daphnia.
Physiological Zoology 58:350 – 355.
Maeda-Martinez, A. M. 1991. Distribution of the species of Anostraca, Notostraca, Spinicaudata, and Laevicaudata in Mexico.
Hydrobiologia 212: 209 – 220.
Maeda-Martinez, A. M., Belk, D., Obregon-Barboza, H., Dumont,
H. J. 1995. Diagnosis and phylogeny of the New World Streptocephalidae (Branchiopoda: Anostraca). Hydrobiologia 298:
15 – 44.
Martin, J. W. 1992. Branchiopoda. Pages 25 – 244 in Harrison, F. W.,
Humes, A. G. Eds. Microscopic Anatomy of Invertebrates. Vol.
9: Crustacea. Wiley – Liss, New York, 625 pp.
Martin, J. W., Felgenhauer, B. E., Abele, L. G. 1986. Redescription of
the clam shrimp Lynceus gracilicornis (Packard) (Branchiopoda,
Conchostraca, Lunceidae) from Florida, with notes on its biology. Zoologica Scripta 15:221 – 232.
Martin, J. W., Belk, D. 1988. Review of the clam shrimp family
Lynceidae Stebbing, 1902 (Branchiopoda: Conchostraca), in the
Americas. Journal of Crustacean Biology 8:451 – 482.
Mathias, P. 1937. Biologie des Crustaces Phyllopodes. Actualites
Scientifiques et Industrielles 447:1 – 107.
McCauley, E. 1984. The estimation of the abundance and biomass of
zooplankton in samples. Pages 228 – 265, in: Downing, J. A.,
Rigler F. H. (Eds.) 1984. A Manual on Methods for the Assessment of Secondary Productivity in Fresh Waters, 2nd edn.
Blackwell Sci., Oxford. 501 pp.
McLaughlin, P. A. 1980. Comparative morphology of recent Crustacea. Freeman, San Francisco, CA.
McQueen, D. J, Post, J. R., Mills, E. L. 1986. Trophic relationships
in freshwater pelagic ecosystems. Canadian Journal of Fisheries
and Aquatic Science. 43:1571 – 1581.
Mellors, W. K. 1975. Selective predation of ephippial Daphnia
and the resistance of ephippial eggs to digestion. Ecology 56:
974 – 980.
Merilainen, J., Huttunen, P., Battarbee, R. W. (Eds.) 1983. Paleolimnology. Proceedings of the Third International Symposium on
Paleolimnology, held at Joensuu, Finland. Hydrobiologia Vol.
103, 318 p.
Merrill, H. B. 1893. The structure and affinities of Bunops
scutifrons, Birge. Transactions of the Wisconsin Academy of
Sciences Arts and Letters. 9:319 – 342.
Michael, R. G., Frey, D. G. 1983. Assumed Amphi-Atlantic distribution of Oxyurella tenuicaudis (Cladocera, Chydoridae) denied by
a new species from North America. Hydrobiologia 106:3 – 35.
Michael, R. G., Frey, D. G. 1984. Separation of Disparalona leei
(Chien, 1970) in North America from D. rostrata (Koch, 1841)
in Europe (Cladocera, Chydoridae). Hydrobiologia 114:81 – 108.
Modlin, R. F. 1982. A comparison of two Eubranchipus species (Crustacea: Anostraca). American Midland Naturalist 107:107 – 113.
Moghraby, A. E. el. 1977. A study on diapause of zooplankton in a
tropical river — the Blue Nile. Freshwater Biology 7:207 – 212.
Monakov, A. V. 1972. Review of studies on feeding of aquatic invertebrates conducted at the Institute of Biology of Inland Waters,
21. Cladocera and Other Branchiopoda
Academy of Science, USSR. Journal of the Fisheries Research
Board of Canada 29:363 – 383.
Moore, M. V., Folt, C. L., Stemberger, R.S. 1996. Consequences of
elevated temperatures for zooplankton assemblages in temperate lakes. Archiv für Hydrobiologie 135:289 – 319.
Moore, W. G., Burn, A. 1968. Lethal oxygen thresholds for certain
temporary pond invertebrates and their applicability to field situations. Ecology 49:349 – 351.
Moore, W. G., Ogren, L. H. 1962. Notes on the breeding behavior of
Eubranchipus holmani (Ryder). Tulane Studies in Zoology
9:315 – 318.
Mort, M. A., Streit, B. 1992. Measuring molecular variation in zooplankton populations: DNA extraction from small Daphnia
species. Aquatic Sciences 54: 77 – 84.
Mossin, J. 1986. Physiochemical factors inducing embryonic development and spring hatching of the European fairy shrimp
Siphonophanes grubei (Dybowsky) (Crustacea: Anostraca).
Journal of Crustacean Biology 6:693 – 704.
Mueller, W. P. 1964. The distribution of cladoceran remains in surficial sediments from three northern Indiana lakes. Invest. Indiana Lakes and Streams 6:1 – 63.
Munuswamy, N., Subramoniam, T. 1985. Influence of mating on ovarian and shell gland activity in a freshwater fairy shrimp Streptocephalus dichotomus (Anostraca). Crustaceana 49:225 – 232.
Nebeker, A. V., Onjukka, S. T., Stevens, D. G., Chapman, G. A.,
Dominguez, S.E. 1992. Environmental Toxicology and Chemistry 11: 373 – 379.
Nilsson, N., Pejler, B. 1973. On the relation between fish fauna and
zooplankton composition in north Swedish lakes. Institute of
Freshwater Research, Drottningholm. Report 53:51 – 77.
Norman, A. M., Brady, G. S. 1867. A monograph of the British Entomostraca belonging to the families Bosminidae, Macrothricidae, and Lynceidae. Natural History Transaction of Northumberland and Durham 1:354 – 408.
North American Lake Management. 1995. 15th International Symposium of the North American Lake Management Society,
Toronto, Ontario, Canada, 6 – 11 November. Lake and Reservoir Management 11:113 – 206.
O’Keefe, T. C., Brewer, M. C., Dodson, S.I. 1998. Swimming behavior of Daphnia: Its role in determining predation risk. Journal
of Plankton Research [in press].
O’Brien. W. J. 1979. The predator – prey interaction of planktivorous
fish and zooplankton. American Scientist 67:572 – 581.
Ojima, Y. 1958. A cytological study on the development and maturation of the parthenogenetic and sexual eggs of Daphnia pulex
(Crustacea – Cladocera). Kwansei Gakuin University Annual
Studies 6:123 – 176, 28 plates.
Olesen, J. 1996. External morphology and phylogenetic significance
of the dorsal / neck organ in the Conchostraca and the head
pores of the cladoceran family Chydoridae (Crustacea, Branchiopoda). Hydrobiologia 330:213 – 226.
Olesen, J. 1998. A phylogenetic analysis of the Conchostraca and
Cladocera (Crustacea, Branchiopoda, Diplostraca) Zoological
Journal of the Linnean Society 122:491 – 536.
Orlova-Bienkowskaja, M. J. 1998. A revision of the cladoceran
genus Simocephalus (Crustacea, Daphniidae). Bulletin of the
Natural History Museum of London (Zoology) 64:1 – 62.
Pace, M. L., Vaque, D. 1994. The importance of Daphnia in determining mortality rates of protozoans and rotifers in lakes. Limnol. Oceanogr. 39:985 – 996.
Persoone, G., Sorgeloos, P. Roels, O., Jaspers, E. (Eds.) 1985. The brine
shrimp Artemia. Vols. 1 – 3. Universa Press: Wettere, Belgium.
Peters, R. H. 1984. Methods for the study of feeding, grazing, and
assimilation by zooplankton. pp. 336 – 412, in: Downing, J. A.,
911
Rigler, F. H. Eds. A Manual on Methods for the Assessment of
Secondary Productivity in Fresh Waters, 2nd edn. Blackwell
Sci., Oxford.
Peters, R. H. 1987. Metabolism in Daphnia. in: Peters, R. H., de
Bernardi, R. Eds. “Daphnia” Memorie dell’Istituto Italiano di
Idrobiologia Vol. 45, pp. 193 – 243.
Porter, K. G. 1973. Selective grazing and differential digestion of
algae by zooplankton. Nature 244:179 – 180.
Porter, K. G. 1977. The plant-animal interface in freshwater ecosystems. American Scientist 65:159 – 170.
Porter, K. G., Feig, Y. S., Vetter, E. F. 1983. Morphology, flow
regimes, and filtering rates of Daphnia, Ceriodaphnia, and
Bosmina fed natural bacteria. Oecologia 58:156 – 163.
Porter, K.G., Gerritsen, J., Orcutt, Jr., J. D. 1982. The effect of food
concentration on swimming patterns, feeding behavior, ingestion, assimilation, and respiration by Daphnia. Limnology and
Oceanography 27:935 – 949.
Porter, K. G., Orcutt, J. D. 1980. Nutritional adequacy, manageability, and toxicity as factors that determine the food quality of
green and blue-green algae for Daphnia. American Society of
Limnology and Oceanography Special Symposium 3:268 – 281.
Potts, W. T. W., Durning, C. T. 1980. Physiological evolution in the
Branchiopods. Comparative Biochemistry and Physiology
67B:475 – 484.
Prepas, E. E. 1984. Some statistical methods for the design of experiments and analysis of samples. Pages 266 – 335, in: Downing,
J. A., Rigler, F. H. Eds. A manual on methods for the assessment
of secondary productivity in fresh waters, 2nd edition. Blackwell Sci., Oxford.
Proctor, V. W. 1964. Viability of crustacean eggs recovered from
ducks. Ecology 45:656 – 658.
Rajapaksa, R. 1986. A contribution to the taxonomy, biogeography
and gamogenesis of freshwater Cladocera, with special reference to the tropical region. Ph.D. dissertation, University of Waterloo, Ontario. xiv, 221, & 21 Appendix pp.
Rajapaksa, R., Fernando, C.H. 1982. The first description of the
male and ephippial female of Dadaya macrops (Daday, 1898)
(Cladocera, Chydoridae), with additional notes on this common
tropical species. Canadian Journal of Zoology 60:1841 – 1850.
Rajapaksa, R., Fernando, C. H. 1987a. Redescription and assignment of Alona globulosa Daday, 1898 to a new genus
Notoalona and a description of Notoalona freyi sp. nov. Hydrobiologia 144:131 – 153.
Rajapaksa, R., Fernando, C. H. 1987b. Redescription of Dunhevedia
serrata Daday, 1898) (Cladocera, Chydoridae) and a description
of Dunhevedia americana sp. nov. from America. Canadian
Journal of Zoology 65:432 – 440.
Richard, J. 1892. Cladoceres nouveaux du Congo. Grimaldina
Brazzai, Guernella Raphaelis, Moinodaphinia Macquerysi.
Mem. Soc. Zool. Fr. 5:213 – 226.
Richman, S. 1958. The transformation of energy by Daphnia pulex.
Ecological Monographs 28:273 – 291.
Richman, S. E., Dodson, S. I. 1983. The effect of food quality on
feeding and respiration by Daphnia and Diaptomus. Limnol.
Oceanogr. 28(5):948 – 956.
Rigler, F. H., Downing, J. A. 1984. The calculation of secondary productivity. Pages 19 – 58, in: Downing, J. A., Rigler, F. H., Eds. A
Manual on Methods for the Assessment of Secondary Productivity in Fresh Waters, 2nd edn. Blackwell Sci., oxford.
Ringelberg, J. 1987. Light induced behavior in Daphnia. Pages
285 – 323, in: Peters, R. H., de Bernardi, R., Eds. “Daphnia.”
Memorie dell’Istituto Italiano di Idrobiologia, Vol. 45.
Rivier, I. K. 1998. The predatory Cladocera and Leptodorida of the
world. Backhuys Publishing, Leiden, The Netherlands. 213 pp.
912
S. I. Dodson and D. G. Frey
Rzòska, J. 1961. Observations on tropical rainpools and general remarks on temporary waters. Hydrobiologia 17:265 – 286.
Sars, G. O. 1900. Description of Iheringula paulensis G. O. Sars, a
new generic type of Macrothricidae from Brazil. Arch. Math.
Naturvidensk. 22(6):1 – 27.
Sars, G. O. 1904. Pacifische Plankton-Crustaceen. (Ergebnisse einer
Reise nach dem Pacific. Schauinsland 1896 / 97). Zoologische
jahrbücher. Abteilung für Systematik, Geographie und Biologie
der Tiere. 19:629 – 646.
Sars, G. O. 1916. The fresh-water Entomostraca of Cape Province
(Union of South Africa). Part I. Cladocera. Annals of the South
African Museums 15:303 – 351.
Sassaman, C. 1995. Sex determination and evolution of unisexuality
in the conchostraca. Hydrobiologia 298:45 – 65.
Schindler, D. 1977. Evolution of phosphorus limitation in lakes. Science 195:260 – 262.
Schmidt-Nielsen, K. S. 1984. Scaling: Why is animal size so important. Cambridge Uni. Press. 241 p.
Schminke, H. K. 1976. The ubiquitous telson and the deceptive
furca. Crustaceana 30: 292 – 300.
Schrimpf, A., Steinberg. C. 1982. Further recordings of the newly observed cladoceran Daphnia parvula, new record in southern
Germany. Archiv für Hydrobiologie 94: 372 – 381.
Schwartz, S. S., Hann, B. J., Hebert, P.D.N. 1983. The feeding
ecology of Hydra and possible implications in the structuring
of pond zooplankton communities. Biological Bulletin 164:
136 – 142.
Schwartz, S. S. 1984. Life history strategies in Daphnia: a review and
predicitons. Oikos 42:114 – 122.
Schwartz, S. S., Hebert, P.D.N. 1987a. Methods for the activation of
the resting eggs of Daphnia. Freshwater Biology 17:373 – 379.
Schwartz, S. S., Hebert, P. D. N. 1987b. Breeding system of Daphniopsis ephemeralis: adaptations to a transient environment.
Hydrobiologia 145:195 – 200.
Schwartz, S. S., Hebert, P. D. N. 1987c. Daphniopsis ephereralis sp.
n. (Cladocera, Daphniidae): a new genus for North America.
Canadian Journal of Zoology 63: 2689 – 2693.
Sergeyev, V. N. 1971. Povedeniye i mekhanizm pitaniya Lathonura
rectirostris (Cladocera, Macrothricidae). Zoologicheskii Zhurnal 50: 1002 – 1010.
Shan, R. K., Frey, D. G. 1983. Pleuroxus denticulatus and P.
procurvus(Cladocera, Chydoridae) in North America: distribution, experimental hybridization, and the possiblity of natural
hybridization. Canadian Journal of Zoology 61:1605 – 1617.
Shapiro, J., Forsberg, B., Lamarra, V., Lynch, M., Smeltzer, E., Zoto,
G. 1982. Experiments and experiences in biomanipulation:
Studies of biological ways to reduce algal abundance and eliminate blue-greens. Interim Report No. 19 of the Limnological
Research Center, University of Minnesota, Minneapolis Minnesota. 251pp.
Shaw, S. R., Stone, S. 1982. Photoreception. Pages 291 – 367 in:
Atwood H.L., Sandeman, D. C., Eds. Neurobiology: Structure
and Function. 479 pp. Vol. III, D. E. Bliss, Editor-in-Chief. The
Biology of Crustacea. Academic Press. New York.
Shei, P., Iwakuma, T., Fujii, K. 1988. Population dynamics of Daphnia
rosea in a small eutrophic pond. Ecological Research 3:291 – 304.
Shen, C.-j., Tai, A.-y., Chiang, S.-c. 1966. (On the cladoceran fauna
of Hsi-song-pang-na and vicinity, Yunnan Province). Acta
Zootaxonomica Sinica 3:29 – 423. [In Chinese, with English
summary.]
Shurin, J. B., Dodson, S.I. 1997. Sublethal toxic effects of cyanobacteria and nonylphenol on environmental sex determination and
development in Daphnia. Environmental Toxicology and Chemistry 16:1269 – 1276.
Sissom, S. L. 1980. An occurrence of Cycloestheria hislopi in North
America. Texas Journal of Science 32:175 – 176.
Smirnov, N. N. 1970. Cladocera (Crustacea) from the Permian of Eastern Kazakhstan. Paleontological Journal 3:1 – 37 [In Russian.]
Smirnov, N. N. 1971. Chydoridae Fauny Mira. Fauna SSSR, Nov.
Ser. No. 101. Rakoobraznyye, T. 1, vyp. 2. 531 p. [Available in
English from Israel Program for Scientific Translations,
Jerusalem 1974.]
Smirnov, N. N. 1985. Anchistropus ominosus sp. n. (Cladocera,
Chydoridae) iz Reki Shingy (pritok Amazonki). Zoologicheskii
Zhurnal 64:137 – 139.
Smirnov, N. N. 1992. The Macrothricidae of the world. Guides to
the identification of the microinvertebrates of the Continental
Waters of the World. SPB Academic Publishing.
Smirnov, N. N. 1996. Cladocera: The Chydorinae and Sayciinae
(Chydoridae) of the world. Guides to the identification of the
microinvertebrates of the continental waters of the world, No.
11. SPB Academic Publishing. The Hague. 197 p.
Smirnov, N. N., Timms, B. V. 1983. A revision of the Australian
Cladocera (Crustacea). Records of Australian Museums, Supplement 1:1 – 132.
Sommer, U. 1989. Plankton ecology: succession in plankton communities. Springer, Berlin.
Spaak, P. 1995. Sexual reproduction in Daphnia: Interspecific differences in a hybrid species complex. Oecologia. 104:501 – 507.
Spitze, K. 1995. The quantitative genetics of zooplankton life histories. Experientia 51:454 – 464.
Sprules, W. G. 1975. Midsummer crustacean zooplankton communities in acid-stressed lakes. Journal of the Fisheries Research
Board of Canada 32:389 – 385.
Sprules, W. G., Riessen, H. P., Jin, E. H. 1990. Dynamics of the
Bythotrephes invasion of the St. Lawrence Great Lakes. Journal
of Great Lakes Research 16:346 – 351.
Starobogatov, Ja.I. 1989. Sistema Crustacea. Zoologicheskii Zhurnal
65:1769 – 1781.
Stirling, G., McQueen, D. J. 1986. The influence of changing temperature on the life history of Daphniopsis ephemeralis. Journal of
Plankton Research 8:583 – 595.
Sublette, J. E., Sublette, M. S. 1967. The limnology of playa lakes on
the Llano Estacado, New Mexico and Texas. The Southwestern
Naturalist 12:369 – 406.
Swanson, G. A., Meyer, M. I., Adomaitis, V. O. 1985. Foods consumed by breeding mallards on wetlands of south-central North
Dakota. Journal of Wildlife Management 49:197 – 203.
Swift, M. C., Federenko, A. Y. 1975. Some aspects of prey capture
by Chaoborus larvae. Limnology and Oceanography 20:
418 – 425.
Tappa, D. W. 1965. The dynamics of the association of six limnetic
species of Daphnia in Aziscoos lake, Maine. Ecological Monographs 35:395 – 423.
Taylor, D. J., Hebert, P.D.N. 1993. Habitat-dependent hybrid parentage and differential introgression between neighboring sympatric Daphnia species. Proceedings of the National Academy
of Sciences of the United States of America 90:7079 – 7083.
Teschner, M. 1995. Effects of salinity on the life history and fitness of
Daphnia magna: Variability within and between populations.
Hydrobiologia 307:33 – 41.
Tessier, A. J. 1983. Coherence and horizontal movements of patch of
Holopedium gibberum (Cladocera). Oecologia 60:71 – 75.
Tessier, A. J. 1986. Comparative population regulation of two planktonic cladocera (Holopedium gibberum and Daphnia catawba).
Ecology 67:285 – 302.
Tessier, A. J., Goulden, C. E. 1982. Estimating food limitation in cladoceran populations. Limnology and Oceanography 27:707 – 717.
21. Cladocera and Other Branchiopoda
Tessier, A. J., Goulden, C. E. 1987. Cladoceran juvenile growth: Implications for competitive ability. Limnology and Oceanography
32:680 – 685.
Thiel, H. 1963. Zur Entwicklung von Triops cancriformis Bosc.
Zoologischer Anzeiger 170:62 – 68.
Thorp, J. H. 1986. Two distinct roles for predators in freshwater assemblages. Oikos 47:75 – 82.
Thorp, J. H., Black, A. R., Haag, K. H., Wehr, J. D. 1994. Zooplankton assemblages in the Ohio River: Seasonal, tributary, and navigational dam effects. Canadian Journal of Fisheries and
Aquatic Sciences 51:1634 – 1643.
Threlkeld, S. T. 1976. Starvation and the size structure of zooplankton communities. Freshwater Biology 6:489 – 496.
Threlkeld, S. A. 1979. Estimating cladoceran birth rates: the importance of egg mortality and the egg age distribution. Limnology
and Oceanography 24:601 – 612.
Threlkeld, S. T. 1986a. Differential temperature sensitivity of two
cladoceran species to resource variation during a blue-green algal bloom. Canadian Journal of Zoology 64:1739 – 1744.
Threlkeld, S. T. 1986b. Life table responses and population dynamics
of four cladoceran zooplankton during a reservoir flood. Journal of Plankton Research 8:639 – 647.
Threlkeld, S. T. 1988. Daphnia population fluctuations: patterns and
mechanisms. Pages 367 – 388 in: Peters, R., deBernardi, R. editors. Daphnia. Memorie Istituto Italiano di Idrobiologia Vol. 45.
Threlkeld, S. T. Willey, R. L. 1993. Colonization, interaction, and organization of cladoceran epibiont communities. Limnology and
Oceanography 38:584 – 591.
Tillmann, U., Lampert, W. 1984. Competitive ability of differently
sized Daphnia species: An experimental test. Journal of Freshwater Ecology 2:311 – 323.
Tollrian, R., Dodson, S.I. 1998. Predator induced defenses in cladocerans. Pages 177 – 202, in: Tollrian, R., Harvell, C. D. Eds. The
ecology and evolution of inducible defenses. Princeton Uni.
Press, Princeton, NJ.
Tollrian, R., Harvell, C. D. 1998. The evolution of inducible defenses: current ideas. in: Tollrian R., Harvell, C. D., Eds. The
ecology and evolution of inducible defenses. Princeton Uni.
Press, Princeton, NJ, pp. 306 – 322
Torrentera, L., Dodson, S. I. 1995. Morphological diversity in populations of Artemia in Yucatan (Branchiopoda). Journal of
Crustacean Biology 15:86 – 102.
Tribbey, B. A. 1965. A field and laboratory study of ecological succession in temporary ponds. Ph.D. Thesis, University of Texas, Austin.
EPA, U. S. 1992. Environmental monitoring and assessment
program: Great Lakes monitoring and research strategy. Office
of Research and Development, Environmental Research
Laboratory-Duluth, Duluth, MN 55804.
Van Der Hoeven, N. 1990. Effect of 3,4-dichloroaniline and metavanadate on Daphnia populations. Ecotoxicology and Environmental Safety 20:53 – 70.
Vanni, M. J. 1986. Competition in zooplankton communities: Suppression of small species by Daphnia pulex. Limnology and
Oceanography 31:1039 – 1056.
913
Walossek, D. 1995. The Upper Cambrian Rehbachiella, its larval
development, morphology, and significance for the phylogeny of
Branchiopoda and Crustacea. Hydrobiologia 298:1 – 13.
Weider, L. J., Hebert, P. D. N. 1987. Ecological and physiological
differentiation among low-arctic clones of Daphnia pulex. Ecology 68:188 – 198.
Weider, L. J., Lampert, W. 1985. Differential response of Daphnia
genotypes to oxygen stress: respiration rates, hemoglobin content and low-oxygen tolerance. Oecologia (Berlin) 65:
487 – 491.
Weismann, A. 1880. Beitrage zur Naturgeschichte der Daphnoiden.
VI. Samen und Begattung der Daphnoiden. Zeitschrift fur wissenschaftliche Zoologie 33:55 – 256.
Wetzel, R. G. 1983. Limnology, 2nd ed., Saunders, Philadelphia, 753 p.
Whiteside, M. C. 1970. Danish chydorid Cladocera: modern ecology
and core studies. Ecological monographs 40:79 – 118.
Whiteside, M. D., Williams, J. B. 1975. A new sampling technique
for aquatic ecologists. International Association of Theoretical
and Applied Limnology. Proceedings. 19: 1534 – 1539.
Whiteside, M. D., Williams, J. B., White, C. P. 1978. Seasonal abundance and pattern of chydorid Cladocera in mud and vegetative
habitats. Ecology 59:1177 – 1188.
Wiggins, G. B., Mackay, R. J., Smith, I. M. 1980. Evolutionary and
ecological strategies of animals in annual temporary pools.
Archiv für Hydrobiologie Supplement. 58:97 – 206.
Williams, G. C. 1975. Sex and evolution. Monographs in Population
Biology No. 8. Princeton, Princeton. 200 pp.
Williams, J. L. 1978. Ilyocryptus gouldeni, a new species of water
flea, and the first American record of I. agilis Kurz (Crustacea:
Cladocera: Macrothricidae). Proceding of the Biological Society
of Washington. 91:666 – 680.
Wiman, F. H. 1979. Mating patterns and speciation in the fairy
shrimp genus Streptocephalus. Evolution 33:172 – 181.
Wiman, F. H. 1981. Mating behavior in the Streptocephalus fairy
shrimps (Crustacea: Anostraca). The Southwestern Naturalist
25:541 – 546.
Wingstrand, K. G. 1978. Comparative spermatology of the Crustacea
Entomostraca. 1. Subclass Branchiopoda. Det Kongelige Danske
Videnskabernes Selskab Biologiske Skrifter 22:1 – 6720pl.
Winkler, D. W. (Ed.) 1977. An ecological study of Mono Lake,
California. Institute of Ecology Publication Number 12, University of California, Davis.
Wolvecamp, H. P., Waterman, T. Respiration. Pages 35 – 100, in:
Waterman, T. Ed. The Physiology of Crustacea. Vol. 1. Academic Press: New York. 670 p.
Zaret, T. M. 1980. Predation and freshwater communities. Yale
University Press. New Haven. Zou, E.M, Fingerman, M. 1997.
Effects of estrogenic xenobiotics on molting of the water flea,
Daphnia magna. Ecotoxicology and Environmental Safety 38:
281 – 285.
Zaffagnini, R., Trentini, R. 1980. The distribution and reproduction
of Triops cancriformis (Bosc) in Europe. Monitore Zoologico
italiano nuova serie 14:1 – 8.
This Page Intentionally Left Blank
22
COPEPODA
Craig E. Williamson
Janet W. Reid
Department of Earth and
Environmental Sciences
Lehigh University
Bethlehem, Pennsylvania 18015
Department of Invertebrate Zoology
National Museum of Natural History
Smithsonian Institution
Washington, DC 20560
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Internal Anatomy and Physiology
III. Ecology and Evolution
A. Reproduction and Life History
B. Distribution
C. Physiological Adaptations
D. Behavioral Ecology
E. Foraging Relationships
F. Population Regulation
G. Functional Role in the Ecosystem
IV. Current and Future Research Problems
V. Collecting and Rearing Techniques
A. Collection and Preservation
B. Rearing Techniques
VI. Identification Techniques
A. Dissection Techniques
for Identification
B. Characteristics for Distinguishing
the Free-Living Freshwater Orders
C. Identification of the Continental
Free-Living Copepoda
Literature Cited
I. INTRODUCTION
The subclass Copepoda H. Milne Edwards, 1840,
in the class Crustacea is the largest of the entomostracan classes with over 10,000 known species. Due
largely to their high densities in the world oceans, they
are estimated to be some of the most abundant metazoans on earth (Hardy, 1970). Copepods are also
abundant in terrestrial systems where they may reach
densities in excess of 100,000 individuals / m2 in wet organic soils (Reid, 1986). Although of marine origin,
copepods may also dominate the fauna of some freshwater systems. For example, in Lake Baikal, which is
both the deepest lake in the world and one which contains 20% of the unfrozen freshwater on earth, a single
species (Epischura baikalensis) contributes up to 96%
of the zooplankton (Huys and Boxshall, 1991).
Free-living freshwater copepods can be distinguished from other small aquatic invertebrates by a variety of morphological characteristics. They have a
somewhat cylindrical, segmented body with numerous
segmented appendages on the head and thorax, and
two setose caudal rami on the posterior end of the abEcology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
domen. They possess an exoskeleton, conspicuous first
antennae, and a single, simple, anterior eye. The defining apomorphy of the Copepoda is the structure of
their swimming legs, each pair of which is connected at
the base by a “coupler” or “intercoxal sclerite”. The
name of the class is from the Greek words kope for oar
and podos for foot, derived from the coupler, which
apparently increases the energetic efficiency of the legs.
Here, we follow Huys and Boxshall (1991) in recognizing two infraclasses and 10 orders of copepods.
The infraclass Progymnoplea includes the order Platycopioida, while the infraclass Neocopepoda contains
the other nine orders in two superorders. The superorder Gymnoplea includes the Calanoida, while the superorder Podoplea includes the remaining eight orders.
Three of the 10 orders (Monstrilloida, Siphonostomatoida, and Poecilostomatoida) are primarily parasitic
on a variety of fish and invertebrates, and the great majority are marine. Some poecilostomatoids, such as
Ergasilus may have free-living planktonic males and
copepodids while the adult females are ectoparasites on
fish. Three other orders (Platycopioida, Misophrioida,
and Mormonilloida) are primarily marine free-living
915
916
C. E. Williamson and J. W. Reid
species of the benthos or plankton. This chapter will
focus on the three orders that contain the widespread,
abundant, and primarily free-living copepods
(Calanoida, Cyclopoida, Harpacticoida), of which
there are over 7750 species. The harpacticoids include
approximately 3000 species, 10% of which inhabit
freshwaters; calanoids include about 2300 species,
25% of which are freshwater; while there are about
2450 species of cyclopoids, 20% of which are freshwater (Bowman and Abele, 1982). Many cyclopoids are
parasitic. The remaining order, Gelyelloida, includes
two recently discovered free-living species in the genus
Gelyella from subterranean habitats in Europe. Another species has been reported from a single location
in North America (Reid, unpublished).
Free-living freshwater copepods generally range in
size from less than 0.5 to 2.0 mm in length, although
some species such as the cyclopoids Macrocylops fuscus
and Megacyclops gigas, and calanoids in several genera
including Heterocope, Epischura, Limnocalanus, and
Hesperodiaptomus can reach lengths of 3 – 5 mm. While
the vast majority of freshwater copepods are transparent or a pale gray or brown, colors may range from
black to red, orange, pink, purple, green and blue. The
brighter colors are often caused by plant pigments such
as carotenoids in oil droplets of the copepod.
Copepods are common in a variety of aquatic and
semiaquatic habitats ranging from moist soils, leafpacks, groundwater, wetlands, and phytotelmata, to
lakes, estuaries, and open oceans. Most species are omnivorous to some extent, with foods ranging from detritus and pollen, to phytoplankton, other invertebrates, and even larval fish. Copepods frequently
comprise a major portion of the consumer biomass in
these habitats. They play a pivotal role in aquatic food
webs both as primary and secondary consumers, and as
a major source of food for many larger invertebrates
and vertebrates.
II. ANATOMY AND PHYSIOLOGY
A. External Morphology
The generalized copepod body in freshwater systems consists of an elongate, segmented body with an
exoskeleton. There is usually a single major articulation
that divides the body functionally into an anterior portion, the prosome, and a posterior portion, the urosome
(Figs. 1 and 2). In the superorder Podoplea which includes the cyclopoids, harpacticoids, and gelyelloids, the
major body articulation (not distinct in the gelyelloids)
is between the fifth and sixth thoracic segments (somites
of the fourth and fifth legs); while in the superorder
Gymnoplea, which includes the calanoids, it is between
the sixth thoracic segment (somite of the fifth leg) and
the genital segment. The head is fused with the first and
sometimes second thoracic segment to form the
cephalosome. The thorax consists of seven segments,
starting with the segment that bears the maxillipeds and
ending with the genital segment. The number of segments in the urosome is variable, but usually between
three and five. The prosome thus includes the head and
most of the thorax, while the urosome includes the last
one or two segments of the thorax and the entire abdomen, except for the terminal caudal rami.
Five pairs of jointed appendages occur on the head:
the first antennae (antennules), second antennae (antennae), mandibles, first maxillae (maxillules), and second
maxillae (Figs. 1 – 3). The first antennae serve many important functions related to reproduction, locomotion,
and feeding. They are armed with both chemoreceptors
and mechanoreceptors that may aid in the discrimination between mates, prey, and potential predators. The
first antennae of male copepods are geniculate (have a
joint that can bend abruptly) and modified for grasping
the female during copulation (Figs. 1 and 3). In male
harpacticoids, cyclopoids, and gelyelloids, both first
antennae are geniculate, while in male calanoids only
the right antenna is geniculate. The second antennae of
calanoids, harpacticoids, and gelyelloids are biramous,
while those of most freshwater cyclopoids lack an exopod and are thus uniramous.
Five or six pairs of well-developed thoracic appendages are generally present. The first thoracic segment bears the maxillipeds, while the second through
the fifth bear four pairs of morphologically similar, biramous swimming legs which are joined by a coupler.
In most freshwater copepods, the somite (or segment)
bearing the first pair of swimming legs is fused with the
head and segment bearing the maxillipeds, forming the
cephalosome (Fig. 2). The swimming legs consist of
two broad basal segments (the coxa, or coxopod and
the basis, or basipod), to which are attached inner (endopod) and outer (exopod) rami of 1 – 3 segments each
(Fig. 3). In cyclopoids and harpacticoids the fifth pair
of legs is highly reduced. In calanoids the fifth legs are
well developed and symmetrical in the female and
highly asymmetrical and modified for grasping females
during copulation in the male (Fig. 3). Cyclopoids and
harpacticoids have vestigial sixth legs, which are larger
in males. In adult gelyelloids the first three or four pairs
of swimming legs have fused coxae and are highly reduced; the fourth legs are absent in the two European
species. The fifth legs are absent, and male gelyelloids
have vestigial sixth legs in the form of flaps that cover
the genital aperture, while females have no sixth legs.
Abdominal appendages are absent in copepods,
22. Copepoda
FIGURE 1
Major body types of freshwater copepods: (A) a calanoid, Arctodiaptomus dorsalis male; (B) a
cyclopoid Mesocyclops americanus female, grasping a chironomid; (C) epibenthic harpacticoids, Attheyella
spinipes, with the male guarding the female; (D) an interstitial harpacticoid, Parastenocaris palmerae, female
with egg sac; (E) an undescribed species of gelyelloid, female; and (F) a first stage nauplius (NI) of
Macrocyclops fuscus [redrawn from Dahms and Fernando, 1994].
917
918
C. E. Williamson and J. W. Reid
FIGURE 2
Generalized body plan of free-living freshwater copepods. Abbreviations: R,
rostrum; A1, first antenna; A2, second antenna; Md, mandible; Mxl, first maxilla, Mx, second
maxilla; Mxp, maxilliped; P1 – P4, first through fourth swimming legs; P5, fifth leg; P6, sixth
leg; GS, genital segment; CR, caudal ramus. The fifth and sixth legs in Gelyelloida, and the
sixth legs in Calanoida, are lacking.
although the abdomen terminates in two caudal rami
that are variously armed with setae and spines.
B. Internal Anatomy and Physiology
The circulatory system of cyclopoids and harpacticoids consists of a hemocoel with no heart or blood
vessels. Blood is circulated by body and gut movements, and respiration occurs through the general body
surfaces. Copepods have no gills or other respiratory
organs. Calanoids have a more developed circulatory
system that includes a dorsal heart with one pair of lateral ostia and one ventral ostium. Blood is carried anteriorly by a short anterior aorta; no other blood vessels
are present.
The digestive system of copepods includes a mouth,
esophagus (foregut), midgut, hindgut, and anus. The
midgut may have a saclike diverticulum or cecum, and
the anus is located at the posterior end of the urosome.
The esophagus and hindgut are chitinized. A variable
number of labral glands are located around the mouth.
They function in a manner analogous to salivary glands
by secreting mucus that mixes with food before entering
the esophagus. The overall structure of the alimentary
canal and the numbers and types of associated cells may
vary with the feeding habits of the species (Musko,
1983). Egested fecal material is surrounded by a peritrophic membrane secreted by cells in the midgut of at
least some calanoid and harpacticoid copepods, forming pellets (Gould, 1957; Fahrenbach, 1962).
Nitrogenous excretion occurs through antennal
glands in naupliar (juvenile) stages, and through maxillary glands in adults. Osmoregulation is accomplished largely by low osmolality of the hemolymph
(200 mOsm, Mantel and Farmer, 1983). Some
harpacticoids have integumental windows on their dorsal cephalothorax that serve as sites of ion exchange,
and aid in osmoregulation (Hosfeld and Schminke,
1997). These windows are fairly common in most freshwater and some estuarine canthocamptids and parastenocaridids, as well as in some species of Halicyclops.
They do not appear in marine copepods.
With the exception of a few species of harpacticoids (see Sarvala, 1979), reproduction in copepods is
sexual. Copulation between male and female followed
by union of haploid gametes is required for the formation of a fertile diploid zygote. Females have a paired
or unpaired ovary dorsal to the midgut, with paired
oviducts extending anteriorly. Diverticuli extend from
the oviducts posteriorly to the gonopore on the genital
segment. In calanoids the diverticuli join just before entering an atrium. Female calanoids also have a pair of
22. Copepoda
FIGURE 3 Copepod body parts and appendages, mainly of a harpacticoid, from anterior to posterior order:
R, rostrum; A1, first antenna; A2, second antenna; Md, mandible; Mxl, first maxilla; Mx, second maxilla;
Mxp, maxilliped; P1 – 4, first through fourth swimming legs; P5, fifth leg; P6, sixth leg; GS, genital segment;
RS, seminal receptacle; and CR, caudal ramus. Other abbreviations: exp, exopod; enp, endopod; and benp,
baseopod. Note differences in the structure of the fifth legs of male and female calanoids (lower right and
lower left in figure) showing the modification of the male fifth leg for grasping the female during mating.
Cyclopoid and harpacticoid fifth legs are smaller, and the basal segment of the harpacticoid P5 is enlarged on
the inner margin (middle left of figure). In the upper right of the figure the second antennae of two different
harpacticoids illustrate the difference in A2 structure with basis and allobasis.
919
920
C. E. Williamson and J. W. Reid
seminal receptacles that open into the atrium. Female
cyclopoids have a single seminal receptacle which is
connected to the gonopores by fertilization ducts.
Glandular cells along the terminal end of the oviducts
secrete material for the egg shells. Skin glands adjacent
to the gonopore secrete the egg-sac membrane.
Male copepods have a single dorsal gonad (testis).
The vas deferens (one in calanoids, two in cyclopoids)
is composed of the seminal vesicle, spermatophoric sac,
and ejaculatory duct. Male calanoids have a single
gonopore, female calanoids and both male and female
cyclopoids and harpacticoids have paired gonopores.
Spermatophores are elongate and cylindrical in
calanoids and harpacticoids, and more compact and often kidney-shaped in cyclopoids (Fig. 4).
The nervous system of copepods is composed of a
supraesophageal ganglion (brain) connected to a ventral nerve cord by two large circumesophageal connectives. The ventral nerve cord has several thoracic ganglia. Copepods have a variety of sensory receptors.
Freshwater copepods have a simple eye spot that cannot form images; they lack the highly developed compound eyes with lenses characteristic of some marine
pontellids (Gophen and Harris, 1981). Most copepods
have a frontal organ consisting of a complex of fine
sensory hairs and pores located on the anterior tip of
their rostrum. The organ, referred to as the sensory
pore X organ, includes the organ of Bellonci (Hosfeld,
1996), is connected to the brain by paired nerves and
probably functions in chemoreception (Elofsson, 1971;
Strickler and Bal, 1973).
The structures of several other types of sensory receptors have been investigated in freshwater copepods,
and their function established by analogy with other
known arthropod receptors. The body surfaces of
calanoids and cyclopoids are covered with two types
of integumental sensilla:small pegs (sensilla basiconica)
and fine hairs (sensilla trichodea). The pegs are thought
to be chemoreceptors, while the hairs may serve as
both chemoreceptors and mechanoreceptors (Strickler,
1975a). Cyclopoids have rows of small spines located
at the joints between the segments of the abdomen, the
swimming legs, and the caudal rami. These spines are
innervated by proprioceptors that sense the position of
the relevant body parts (Strickler, 1975b). The first antennae of many cyclopoids and calanoids have setae
that are modified ciliary processes that may serve in
mechanoreception (Strickler and Bal, 1973; Friedman,
1980). The mouthparts of diaptomids have numerous
contact chemoreceptors, but no mechanoreceptors
(Friedman and Strickler, 1975; Friedman, 1980). Experiments in which copepods were fed flavored and unflavored polystyrene beads confirm the chemoreceptive
abilities of copepods (DeMott, 1986).
FIGURE 4
Genital segments with attached spermatophores on
females of (A) a calanoid, Arctodiaptomus dorsalis, (B) a harpacticoid, Attheyella spinipes, and (C) a cyclopoid, Metacyclops cushae.
Calanoids may bear one to several spermatophores, while harpacticoids usually bear only one, and cyclopoids always bear two.
III. ECOLOGY AND EVOLUTION
A. Reproduction and Life History
Copepods develop from fertilized eggs which hatch
into a larval stage called a nauplius (Fig. 1F). There are
six naupliar stages (N1 – N6) followed by six copepo-
22. Copepoda
did stages (C1 – C6), the last of which is the adult.
[Some authors use the term “copepodite” instead of
copepodid, but the “ite” ending is more properly used
to refer to a part of a structure.] Growth is determinate, and adults do not continue to molt as in cladocerans. The nauplius larva starts out with a somewhat
rounded body with three pairs of appendages: the first
antennae, the second antennae, and the mandibles.
During development the body of the nauplius becomes
more elongate, additional appendages appear, and existing appendages become more elongate and specialized. A pronounced metamorphosis occurs between the
last naupliar and the first copepodid instar. Copepodids
are morphologically much more similar to the adults,
as the body is clearly divided into a prosome and a
urosome with caudal rami (Fig. 5).
The sexes of adult copepods are dimorphic. The
mechanism of sex determination is genetic in some
species but not known in others (Wyngaard and Chinappa, 1982). Sexual dimorphism is characterized by
differences in the structure of the first antennae and the
fifth and sixth legs, as well as by the number of urosomal segments and the generally larger size of the females versus the males (see taxonomy section). Sexual
dimorphism in body size is more pronounced in the
cyclopoids than in the calanoids or harpacticoids. Cyclopoids have a mean female:male body length ratio of
1.44; while for calanoids, this ratio is 1.13 (Gilbert and
Williamson, 1983). In some cyclopoids the third and
fourth swimming legs are dimorphic, while sexual dimorphism of the swimming legs is most pronounced in
harpacticoids where some or all of the five pairs of
swimming legs may be dimorphic.
Morphological variation brought about by allometric growth of certain body parts (cyclomorphosis) is
subtle and not well understood in copepods. Seasonal
variation in adult body size is common, with body size
generally being smaller during the warmer summer
months; both temperature and food may play a role in
this seasonal variation (Reed, 1986; Reed and Aronson, 1989; Lescher-Moutoué, 1996). Some cyclopoids
also may have more plumose setae on their caudal rami
and swimming legs during the summer (LescherMoutoué, 1996). In a study where 22 different body
dimensions were measured, allometric variation was established in Tropocyclops prasinus (Riera and Estrada,
1985).
The development time from extrusion to hatching
for subitaneous (nondiapausing) eggs is usually between 1 and 5 days. Development time from egg to
adult is normally on the order of 1 – 3 weeks, while the
adult life span is generally from one to several months.
Development times and life spans are much longer at
lower temperatures and may vary considerably with
921
species. Some cyclopoids have periods of diapause that
may extend their life spans up to 1 – 2 years. Males
generally mature more rapidly and have shorter life
spans than females.
After reaching maturity copepods begin to produce
gametes and can mate and produce viable embryos
(usually called eggs) within a few days of maturation.
Under favorable conditions, a single adult female produces a new clutch of eggs every few days and several
hundred eggs over her lifetime. Clutches of 2 – 50 or
more eggs may be carried laterally in two egg sacs (cyclopoids), ventrally in a single egg sac (most calanoids
and harpacticoids), or scattered freely in the water (certain calanoids such as Limnocalanus and Senecella).
Diel periodicities in egg laying have been reported for
Mesocyclops ogunnus (Gophen, 1978; the original
species determination of M. leuckarti was later corrected to M. ogunnus).
Calanoids and harpacticoids may produce either of
two types of eggs. The normal mode of reproduction is
through subitaneous eggs which hatch within a few
days of being extruded. Resting eggs (diapausing eggs)
that are capable of extended periods of dormancy may
also be produced in many freshwater as well as coastal
marine calanoids (Hairston, 1996; Hairston and Van
Brunt, 1994). Initial studies on the “egg banks” of diaptomid copepods suggested that these resting eggs
could survive several years (De Stasio, 1990), but more
recent studies have demonstrated that mean ages of the
resting eggs in these egg banks may be several decades,
and that some eggs may even remain viable for up to
several hundred years (Hairston et al., 1995). Some
calanoids, such as Limnocalanus macrurus, may produce only resting eggs in some systems but both resting
eggs and subitaneous eggs in other systems (Roff,
1972). Cyclopoids produce only subitaneous eggs, but
may enter diapause as copepodid stage II — adult, including fertilized females (Dobrzykowski and Wyngaard, 1993; Naess and Nilssen, 1991). These resting
stages may vary from a simple developmental arrest to
a fully encysted diapausing stage. In temporary ponds,
cyclopoids such as Diacyclops can emerge from diapause within a single day of the appearance of water
(Wyngaard et al., 1991). Cyclopoid diapause is considered a true diapause similar to that observed in insects
(Elgmork and Nilssen, 1978; Naess and Nilssen,
1991). A few species of harpacticoids may encyst in response to adverse environmental conditions (Sarvala,
1979; Frenzel, 1980).
Diapause can be initiated during either the spring
and summer or the autumn. The proximate environmental cues that trigger the onset of diapause stages (eggs or
copepodids) include overcrowding, photoperiod, and
possibly age (Walton, 1985). The ultimate factors that
922
C. E. Williamson and J. W. Reid
FIGURE 5
Copepodid and adult stages of calanoid (upper) and cyclopoid (lower) copepods. Secondary sex
characteristics are apparent in the preadult stage (fifth copepodid, CV).
22. Copepoda
confer a selective advantage on diapausing individuals
may be related to the avoidance of adverse environmental conditions, including an increase in predation pressures (Hairston and Olds, 1984; Hairston and Walton,
1986). The amphipod Gammarus lacustris preys on
copepod resting eggs in the sediments and may deplete
the “egg bank” to the extent that later recovery from adverse conditions such as the presence of fish predators is
not possible (Parker et al., 1996). The termination of diapause may be caused by either endogenous mechanisms,
exogenous factors (such as low oxygen concentrations,
changes in temperature, light, or physical disturbance),
or some combination of these (Elgmork, 1967).
Female cyclopoids and harpacticoids can store
sperm and thus produce fertile clutches of eggs for extended periods of time in the absence of males (Whitehouse and Lewis, 1973; Hicks and Coull, 1983). Most
calanoid copepods, on the other hand, must mate repeatedly in order to continue clutch production. Diaptomids must mate before each clutch is produced (Watras and Haney, 1980; Williamson and Butler, 1987),
while Epischura can produce multiple clutches from a
single mating (Chow-Fraser and Maly, 1988). Under
favorable environmental conditions female diaptomids
alternate between gravid (oviducts filled with mature
ova) and nongravid reproductive phases. Females must
mate when they are gravid in order to produce viable
eggs (Watras and Haney, 1980; Watras, 1983a, b;
Buskey, 1998; Lonsdale et al., 1998).
Mating behavior is similar for freshwater and marine calanoids (Blades and Youngbluth, 1980; Watras,
1983a; Lonsdale et al., 1998). However, while male marine calanoids can locate females at distances of up to 20
cm with the aid of sex pheromones, diaptomids seem to
respond to mates at distances of only a few body lengths
(Watras, 1983a). Evidence for sex pheromones in freshwater calanoids is scarce (van Leeuwen and Maly,
1991). Watras (1983a) has described a behavioral sequence for mating in diaptomids wherein the male actively pursues the female (pursuit), grasps the female
with his geniculate first antenna (precopula 1), grasps
the female’s urosome with his chelate right fifth leg (precopula 2), and uses the left fifth leg to attach a cigarshaped spermatophore to the genital segment of the female (copula). The spermatophore is affixed with the aid
of a cementlike secretion. During both precopula 2 and
copula, the male and female are aligned with their urosomes adjacent and their metasomes extending in opposite directions. It is likely that precopula is accompanied
by stroking behavior, as occurs in marine calanoids
(Blades and Youngbluth, 1980). The entire mating sequence lasts from one to several minutes. Variations in
sexual size dimorphism in calanoids may be important
to mating success in some species, but not in others
923
(Grad and Maly, 1988, 1992). Eggs are fertilized upon
extrusion from the genital segment. In diaptomids, eggs
that are not fertilized are released into the environment
and disintegrate rapidly (Watras and Haney, 1980;
Williamson and Butler, 1987).
Mating behavior is similar in harpacticoid and cyclopoid copepods, but the behaviors of both groups
differ somewhat from the reproductive activities of
calanoids (Gophen, 1979; Glatzel and Schminke, 1996;
Dürbaum, 1995, 1997). Male harpacticoids and cyclopoids are more active swimmers than females, and
mates are encountered randomly, with no evidence of
sex pheromones. When encountering a female, a male
grasps her with both his geniculate antennae; usually
her third or fourth legs or urosome are held, but any
part of her body will do. A period of high activity follows, during which the pair may jump and turn
through the water. While still grasping the female with
his first antennae, the male works himself into the final
mating position with his ventral metasome facing the
ventral surface of the female’s genital segment so that
the two bodies are parallel but facing in opposite directions. The male strokes this area of the female with his
swimming legs and, with sharp flexing of the abdomen,
deposits a compact spermatophore on her genital segment. Both individuals may remain passive for 5 – 10 s
after spermatophore transfer. The entire mating process
usually takes from 5 to 20 min (in some cases 2 h or
more), after which the male releases the female. In
some harpacticoids the male may retain his grasp on
the female for up to several hours or even days in what
is known as postcopulatory mate guarding (Dürbaum,
1995). During this period the female swims around
with the passively attached male presumably preventing access to the female by any other males. Fertilization occurs upon extrusion of the eggs.
In some harpacticoids precopulatory mate-guarding
has also been observed. This involves the male attaching
to copepodid stage females (as early as CI in some
species) so that they maximize their chances of being able
to mate with a virgin female (Dürbaum, 1997). Interspecific mating has been observed in some cyclopoid species
(Maier, 1995), and hybridization has been reported between two species of Leptodiaptomus (Chen et al., 1997).
Feifarek et al. (1983) have shown that there may
be a cost to mating for cyclopoid copepods. In a laboratory experiment, female Mesocyclops edax that
mated and reproduced throughout their lives exhibited
lower survivorship than unmated females. Within the
group of females which mated, however, there was no
relationship between reproductive output and survival.
This suggests that the cost is associated with the mating
itself rather than with allocation of resources to reproduction versus survival.
924
C. E. Williamson and J. W. Reid
Environmental factors have a great influence on
the life-history characteristics of copepods. Temperature, food availability, and predation are the three factors most often implicated as causes of variations in
life-history traits. Adult female body size and interclutch duration are generally inversely related to temperature, while clutch size shows no consistent relationship to temperature (but see Chow-Fraser and
Maly, 1991). Clutch size is often positively correlated
with female body size (Maly, 1973; Hebert, 1985;
Chow-Fraser and Maly, 1991). Food availability has
also been demonstrated to be an important factor limiting rate of egg production and body size both in the
laboratory and in the field (Edmondson, 1964; Elmore,
1983; Jersabek and Schabetsberger, 1995). Some
species of diaptomids exhibit high reproduction during
winter algal blooms (Vanderploeg et al., 1992a). In
some marine copepods ingestion of certain species of
dinoflagellates can cause production of low-quality
sperm that results in poor fertilization and hatching
failure of eggs (Ianora et al., 1999).
Food limitation can decrease fecundity by either
decreasing clutch size, increasing interclutch duration,
or both. The highly transparent bodies of many copepods allow one to monitor the presence of ova in the
oviducts as well as external eggs carried by females.
The proportion of a copepod population that occurs in
each of these reproductive phases at one point in time
can be used to quantify the relative importance of food
and (in calanoids) mate limitation in nature (Fig. 6;
Williamson and Butler, 1987). The presence or absence
of ova in the oviducts alone are not by themselves,
however, a good indicator of reproductive activity
(Watras and Haney, 1980). This may be due at least in
part to mate limitation and to an increase in the per-
FIGURE 6 Four major reproductive phases of clutch-bearing
female diaptomid copepods: G, gravid; O, ovigerous; N, neither; B,
both. When abiotic environmental conditions are favorable for
reproduction, transitions between phases are influenced by the
availability of males and food. Phase B may be absent in species
where embryos are released before oocyte maturation. [From
Williamson and Butler, 1987.]
centage of gravid individuals caused by the need for diaptomids to re-mate before each clutch is produced.
Predation may influence adult body size (Carter
et al., 1983), sex ratio (Hairston et al., 1983; Svensson,
1997), body pigmentation, or the timing of diapause
events in copepods (Hairston and Olds, 1984; Hairston
and Walton, 1986). Some of these life history variations have a definite genetic component to them and,
therefore, represent evolutionary adjustments rather
than just simple acclimatory responses (Hairston and
Walton, 1986; Wyngaard, 1986). When the invertebrate predator Chaoborus preys upon clutch-carrying
diaptomids, the clutches may become detached and
thus not be ingested; larger clutches are spared more
often than smaller clutches (Svensson, 1996).
Life-history events of copepods may also differ
among populations. Florida populations of Mesocyclops edax have shorter maturation times than populations of conspecifics in Michigan (Wyngaard, 1986).
These differences seem to be largely genotypic, as evidenced by their persistence through two generations in
the laboratory under the same controlled environmental conditions, as well as their persistence during a reciprocal transfer experiment carried out with the two different populations (Wyngaard, 1998).
B. Distribution
Copepods are found in a wide variety of aquatic
environments, ranging from the benthic, littoral, and
pelagic waters of lakes and oceans, to swamps, wetlands, marshes, large rivers, temporary ponds, and
small puddles. Numerous species are common in interstitial and subterranean systems as well. Copepods are
often common in phytotelmata, the small volumes of
water that collect in the various parts of plants (Janetzky et al., 1996; Reid and Janetzky, 1996). Copepods
are present but less abundant in the flowing waters of
streams and rivers. Although most prevalent in aquatic
habitats, copepods have been collected from many
semiterrestrial habitats such as moist forest soil, leaf litter, and even arboreal mosses (Reid, 1986). Calanoids
are primarily planktonic, while cyclopoids and
harpacticoids are generally associated with substrates
in littoral or benthic habitats, although a few species of
cyclopoids are planktonic and may contribute substantially to the zooplankton biomass in many lakes and
ponds. Benthic copepods generally do not have pelagic
larvae, and dispersal may occur in all developmental
stages (Hicks and Coull, 1983).
The copepod component of the zooplankton is
usually dominated by one or two species of calanoid or
cyclopoid copepods. The most common and widely distributed planktonic cyclopoid species in North America
22. Copepoda
are probably Mesocyclops edax, Diacyclops thomasi,
and Acanthocyclops spp. Species richness in benthic
copepods generally shows little or no clear pattern with
latitude in the Americas, but the number of genera of
Canthocamptidae correlates with latitude (Reid, 1992).
The distribution of many copepods seems to be related to temperature. For example, Mesocyclops often
dominates in the south and in the warmer months;
while in the north and in the cooler months in intermediate climates, Mesocyclops is often replaced by either
Cyclops, Diacyclops, or Acanthocyclops. In systems
where Mesocyclops does not reach high densities (e.g.,
Lake Michigan), these other cyclopoids may persist
through the summer. The only calanoid genus endemic
to North America is Osphranticum. The distribution of
planktonic copepods in the northeastern United States
indicates that historically some species depended on
surface waters along retreating glacial ice fronts for
postglacial dispersal; while other species, commonly
found in smaller ponds and at high elevations less accessible to glaciers, may have been aided in their dispersal by their capability for diapause (Stemberger,
1995). Glaciation has strong residual effects on the present-day distribution of hyporheic (streambed) cyclopoids. Previously glaciated sites contain fewer
species of interstitial specialists and fewer narrowly endemic species, but generalist species have apparently
easily reinvaded these areas (Strayer and Reid, 1999).
The pH of a system may also influence the distribution and abundance of copepods, but the effects vary
with species. In a study of 20 small lakes in the northeastern United States ranging in pH from 4.5 to 7.2,
Mesocyclops edax and Leptodiaptomus minutus were
both observed across the full range of pH values, but
the former was more abundant at higher pH values
while the latter was more abundant at lower pH values
(Confer et al., 1983). In this same study Epischura lacustris and Cyclops scutifer were found only at pH values of 5.3 or above. Bioassay experiments showed
100% mortality within 8.5 h for Mesocyclops leuckarti
at pH values of 5, or of 9.2 (Ramalingam and
Raghunathan, 1982). The distribution and abundance
of various copepod species may also vary with lake
size, depth, water transparency, color, and the presence
of vertebrate and invertebrate predators. The horizontal distribution of copepods may be influenced by
aquatic macrophytes (Gehrs, 1974) or fish (Carter and
Goudie, 1986).
In benthic habitats, copepods occur primarily in
the top 1 – 2 cm of the sediments, although diapausing
animals may be found up to depths of 10 – 20 cm or
more. The species richness of copepods in lake sediments is usually between 7 and 25 species, with about
equal numbers of harpacticoids and cyclopoids
925
(Strayer, 1985). The Laurentian Great Lakes together
harbor 30, mostly benthic species of cyclopoids and 34
harpacticoid species (Hudson et al., 1998). The vertical
distribution of copepods within the sediments is primarily dependent upon redox potential, and only a few
species can survive anaerobic sediments (Hicks and
Coull, 1983).
The abundance of harpacticoids in benthic communities tends to increase with larger sediment particle
size (Hicks and Coull, 1983). The most common
species of harpacticoids are in the cosmopolitan family
Canthocamptidae, with the most common North
American genera being Attheyella, Bryocamptus, and
Canthocamptus. Harpacticoids in the family Parastenocarididae are also abundant in well-oxygenated
groundwater and interstitial environments. Common
littoral – benthic cyclopoids include Acanthocyclops,
Diacyclops, Eucyclops, Macrocyclops, and Paracyclops.
C. Physiological Adaptations
Copepods face a wide range of fluctuating environmental conditions in littoral, benthic, pelagic, and
semiterrestrial habitats that require physiological adaptation. Perhaps the most widespread physiological
adaptation of copepods is their ability to respond to
seasonally unfavorable environmental conditions by reducing their metabolic rate and entering diapause in either the egg or late copepodid stages (see Section IIIA).
Diapause may be induced by changes in temperature
(Smyly, 1962) or oxygen concentration (Wierzbicka,
1962). These resting stages are highly resistant to temperature extremes and desiccation.
Physiological responses to temperature may vary
among species and even sex of the copepod. Surfacedwelling harpacticoid species that are exposed to a
greater seasonal variability in food availability and
temperature can also better regulate respiration rates
with temperature than can species living deeper in the
sediments (Gee and Warwick, 1984). The longevity of
adult male Megacyclops viridis may be extended under
fluctuating versus constant temperature regimes,
whereas female longevity is reduced under similar conditions (Smyly, 1980).
Copepods are frequently found in the benthic sediments of productive lakes and ponds which become
hypoxic or anoxic. While low oxygen concentrations
may induce diapause, many copepods still remain active under these rigorous oxygen conditions. A few
planktonic species (Mesocyclops edax, Microcyclops
varicans) vertically migrate into anoxic hypolimnetic
waters for several hours on a daily basis (Williamson
and Magnien, 1982; Threlkeld and Dirnbirger, 1986).
926
C. E. Williamson and J. W. Reid
Laboratory experiments have demonstrated that adult
M. edax can tolerate anoxia for 6 – 12 h at 24.6°C
(Woodmansee and Grantham, 1961), while adult M.
varicans live for at least 36 h. Cyclopoid nauplii can
survive anoxia for less than 1 h at unspecified temperatures (Chaston, 1969). Benthic cyclopoids may persist
for at least four days, and probably indefinitely under
hypoxic (25% saturation, 8°C) conditions, but die
within 2 – 5 h under anoxic conditions at 10°C (Tinson
and Laybourn-Parry, 1985). Females are more tolerant
of anoxia than males, and smaller species resist anoxia
longer than larger species. A few species of harpacticoids are also known to tolerate anoxia (Hicks and
Coull, 1983).
In surface waters where oxygen is more plentiful,
solar radiation may have lethal effects on copepods
(Hairston, 1976; Ringelberg et al., 1984; Williamson et
al., 1994). The presence of plant-derived carotenoid
pigments such as astaxanthin in the body, low temperature conditions, or nutritional state may reduce these
lethal light effects (Hairston, 1976, 1979a – c; Ringelberg, 1980; Ringelberg et al., 1984; Luecke and
O’Brien, 1981). The darker color of the pigmented
copepods may, however, make them more vulnerable to
visually feeding vertebrate predators. Some species
seem to have overcome these conflicting selective pressures by producing two separate morphs: one which
has dark red pigments that increase survivorship under
high light conditions, and one with a carotenoid – protein complex that is a pale green in systems where vertebrate predation pressures are high (Hairston, 1976;
Luecke and O’Brien, 1981).
D. Behavioral Ecology
Copepods are active and highly proficient swimmers that exhibit a diversity of behavioral responses to
environmental stimuli, ranging from the intensity and
wavelength of light to the prospective reward or danger
of other approaching organisms. The swimming mode
of copepods varies with the species and habitat. Benthic harpacticoids and cyclopoids swim and crawl over
substrates in the littoral, benthic, and interstitial habitats. The swimming behavior of planktonic copepods
has been investigated in some detail (Rosenthal, 1972;
Strickler, 1982) and has been reviewed for cyclopoids
(Williamson, 1986).
Planktonic cyclopoids use their swimming legs,
first antennae, and urosome to swim through the water,
alternating between an active hop and a passive sink
phase. The hop phase lasts about 20 ms and consists of
a posteriorly directed thrust of the first antennae and a
metachronal (sequential, one pair at a time, back to
front) thrust of each of the pairs of swimming legs. The
hops occur about once every second and during normal
swimming will accelerate the copepod to a maximum
velocity of about 80 mm / s. The sinking phase is the
time between hops during which the copepod remains
passive.
Many variations on this generalized swimming behavior occur. For example, the hops may have a reduced intensity and increased frequency and appear to
be more of a shuttle behavior than a hop and sink. In
addition, most cyclopoids swim right-side up, while
others such as the small, more herbivorous Tropocyclops, swim upside down. When startled, cyclopoids
may exhibit escape responses that consist of continuous, more vigorous hops which may accelerate the
copepod up to a velocity of 350 mm / s. Cyclops singularis adults, which are on the order of 2 mm in body
length, can move over 10 cm in a single jump — a behavior so pronounced that it allows this copepod to be
separated from other species (Einsle, 1996b).
The swimming behavior of calanoids is distinctly
different from that of cyclopoids. Calanoids spend a
majority of their time gliding slowly through the water,
propelled by the rapid (20 – 80 Hz) vibration of their
feeding appendages. The more carnivorous species,
such as Epischura and Limnocalanus, may actively
cruise through the water in an upright position (dorsal
surface up), while the more herbivorous species tend to
hang vertically or at a slight angle in the water with
their anterior end up. Every few seconds the gliding
motion may be interrupted by a hop that involves the
same appendages and motions as observed in cyclopoids. Less frequently, calanoids stop vibrating their
appendages and sink passively through the water.
Nauplii spend a larger proportion of their time in a
stationary position than do adults and copepodids;
calanoid nauplii vibrate their appendages for only
short periods between passive phases, while cyclopoid
nauplii hop less frequently than do copepodids or
adults (Gerritsen, 1978). Nauplii sink very slowly, if at
all, but can exhibit escape responses which are the
fastest recorded for any metazoan relative to body size
(364 body lengths / s, 55 mm / s, Williamson and
Vanderploeg, 1988).
Copepods alter their swimming behavior in response to many diverse environmental stimuli, including the presence of food (Williamson, 1981) and predators (De Stasio, 1993; Ramcharan and Sprules, 1991).
Light seems to be a particularly important stimulus in
regulating the distribution of copepods. For example,
Heterocope septentrionalis exhibits a swarming behavior in which it preferentially aggregates over light-colored substrates (Hebert et al., 1980), and Leptodiaptomus tyrrelli may swarm to densities exceeding 11,000
per liter in response to contrasting substrates (Byron et
22. Copepoda
al., 1983). Copepods may also respond to different
wavelengths of light in different ways. For example,
Hesperodiaptomus nevadensis swims faster in blue
light than in red, and individuals that contain large
amounts of red carotenoid pigments swim faster than
those that do not (Hairston, 1976). Light is also responsible for the distinct “avoidance of shore” (“Uferflucht”) exhibited by pelagic copepods (Siebeck, 1969,
1980).
The diel vertical migration of planktonic copepods
through the water column seems to be driven largely by
responses to light, predation, and food limitation
(Hutchinson, 1967; Williamson and Magnien, 1982;
Neill, 1990). The most commonly observed migration
patterns consist of downward migrations of several meters into the deeper, darker strata at dawn, and a similar distance upward into the surface waters at dusk.
Reverse migrations downward at dusk have also been
noted, particularly in response to invertebrate predation (Neill, 1990; Neill, 1992). Adult females often migrate farther than do either males or juveniles (Burgi et
al., 1993; Williamson and Magnien, 1982).
The adaptive significance of these vertical migrations is thought to be related to the reduction of predation pressures by predators (Zaret and Suffern, 1976;
Wright et al., 1980; Neill, 1990, 1992), but vertical migration may also influence food availability to the
predators (Williamson et al., 1989; Williamson, 1993;
Makino and Ban, 1998). Changes in vertical overlap
between predatory copepods and their prey populations can be quantified with a predation risk model
that couples behavioral responses with field distribution and migration patterns of predator and prey
(Williamson, 1993). The damaging effects of visible radiation cannot, however, be ruled out as an additional
ultimate cause of these migrations. Damaging solar ultraviolet radiation may penetrate to depths of 10 m or
more in clear lakes with low dissolved organic carbon
concentrations (Morris et al., 1995; Williamson et al.,
1996) and thus can cause substantial mortality in copepods that remain in the surface waters throughout the
day (Williamson et al., 1994). Interestingly, sublethal
concentrations of free cupric ions can reduce the phototactic sensitivity of some estuarine and nearshore marine copepods such as Acartia and Temora (Stearns and
Sharp, 1994).
Copepods may also respond to changes in the density of food in the environment. Cyclopoid copepods
exhibit two distinct types of looping behavior: horizontal and vertical looping (Williamson, 1981). Horizontal
loops consist of the copepod swimming in horizontal
circles in a normal hop and sink swimming mode.
When food densities are high, the frequency of horizontal looping behavior increases and serves to main-
927
tain the copepod in the vicinity of high food densities.
Predatory cyclopoids may swim in rapid, tight, vertical
loops after contacting individual prey organisms. This
behavior aids the copepods in capturing prey and in recovering prey that attempt to escape after an initial attack (Williamson, 1981). Calanoids also alter their
swimming behavior in response to the presence of
food. Senecella calanoides and Epischura lacustris both
increase the frequency of their passive sinking phase in
the presence of prey. This behavior may reduce the
“noise” levels of the predators and thereby permit
them to approach and capture their prey by surprise
(Wong and Sprules, 1986).
In addition to behavioral responses to food, copepods may also react to predators by altering their behavior. Leptodiaptomus tyrrelli reduces its filtering rate
(Folt and Goldman, 1981), and Leptodiaptomus minutus decreases the frequency of its hops (Wong et al.,
1986) in the presence of predatory copepods. Similarly,
Cyclops vicinus reduces its activity levels when the vertebrate predator Abramis brama (bream) is near (Winfield and Townsend, 1983). These diminished activity
levels decrease the ability of both types of predators to
detect these small copepods.
E. Foraging Relationships
The foraging relationships of copepods have received a great deal of attention over the years because
of the important intermediate position of copepods between the phytoplankton and fish in aquatic food
chains, and the fact that copepods are so abundant in a
wide variety of aquatic ecosystems. Copepods contribute a major portion of the biomass and secondary
production in a wide variety of aquatic communities.
The importance of copepods in these communities derives largely from their impact on energy and material
processing, prey mortality rates, and depression of
available resources resulting from foraging relationships.
1. Diet
Copepods feed on a wide variety of foods including algae, pollen, detritus, bacteria, rotifers, crustaceans, dipteran larvae including mosquitos, chironomids, and chaoborids, and even larval fish (Fryer,
1957a, b; Monakov, 1976; Fischer and Moore, 1993;
Hartig et al., 1982; Reid, 1989b; Brandl, 1998). Therefore, they occupy three of the four major trophic positions in the food chain: detritivore; herbivore; and carnivore. Most species of calanoids and cyclopoids are
omnivorous but selective in their feeding (Adrian,
1991; Williamson and Butler, 1986). Calanoids ingest
particles ranging in size from small (a few micrometers)
928
C. E. Williamson and J. W. Reid
algae and bacteria to larger algae, microzooplankton,
and macrozooplankton (1 mm). Cyclopoids are
grasping feeders that generally eat larger foods than
calanoids. Their diet may include algae, rotifers, copepods, other crustaceans, oligochaetes, chironomids, nematodes, larval fish, and a variety of other foods. This
creates an interesting situation where cyclopoids can
prey on the larvae of their own predators, and may
cause significant declines in the predator populations
(Fischer and Frost, 1997; Fischer and Moore, 1993).
Smaller cyclopoids such as Tropocyclops prasinus mexicanus tend to be more herbivorous than larger cyclopoid species (Adrian and Frost, 1992). Gelatinous
chlorophytes are usually considered inedible or of low
food value for zooplankton, but they can support survival and reproduction in some diaptomids (Stutzman,
1995).
Harpacticoids consume microalgae, fungi, protozoa, bacteria, and detritus, and some take larger prey
such as nematodes. The algae may include phytoflagellates, cyanobacteria, and both epipelic and epiphytic
diatoms. Harpacticoids also appear to use dissolved organic matter as a food source (Hicks and Coull, 1983).
The microbial communities associated with dead foods
may be more important than the foods themselves in
the nutrition of harpacticoids. Although carnivory has
been reported in harpacticoids, predation seems to be
relatively unimportant as a mechanism for obtaining
food (Hicks and Coull, 1983). However, Phyllognathopus viguieri is an active predator on soil and plant parasitic nematodes (Lehman and Reid, 1993).
The structure of the feeding appendages often correlates with dietary preferences, and both chemoreception and mechanoreception are important in selection
of different types of food (Price et al., 1983; DeMott,
1986; Legier-Visser et al., 1986). Diaptomids are highly
selective in their feeding and tend to prefer larger, more
nutritious algae, or even small invertebrates such as rotifers (Williamson and Butler, 1986; DeMott, 1995a;
Vanderploeg, 1994). Selectivity in diaptomids varies
with food concentration in a way that is consistent
with optimal foraging models (DeMott, 1995b). While
diaptomids are capable of ingesting filamentous and
colonial cyanobacteria at substantial rates (Schaffner et
al., 1994), they exhibit a distinct avoidance of toxic
strains (DeMott and Moxter, 1991); existing data suggest that diaptomids are more sensitive than Daphnia
to cyanobacterial toxicity (DeMott et al., 1991). When
feeding on filamentous diatoms or cyanobacteria, diaptomids may bite the ends off, a phenomenon known as
“filament clipping” which may in turn reduce filament
length in natural phytoplankton populations (DeMott
and Moxter, 1991; Schaffner et al., 1994; Vanderploeg
et al., 1988).
Both calanoid and cyclopoid copepods feed on ciliates, although many ciliates have behavioral, morphological, or chemical defenses that reduce their vulnerability to attack, capture, or ingestion (Williamson,
1980; Burns and Gilbert, 1993; Hartmann et al., 1993;
Wickham, 1995; Burns and Schallenberg, 1996). Reproduction is enhanced in diaptomids when ciliates are
added to algal foods, indicating that the ciliate biomass
is assimilated and utilized rather than just ingested
(Sanders et al., 1996). Thus diaptomids, which were
once thought to use only algal biomass, can also use
carbon sources from larger heterotrophs such as rotifers, and from protozoan carbon derived from the microbial component of the foodweb.
Diet may vary with the developmental stage, especially in the more carnivorous species. Nauplii and
early copepodids are generally more herbivorous. The
transition to more predatory feeding occurs in the late
copepodid stages in calanoids (Maly and Maly, 1974;
Chow-Fraser and Wong, 1986), while in cyclopoids
even the earlier copepodid stages may be predatory
(Brandl and Fernando, 1986; Williamson, 1986). Adult
females are generally larger in size and correspondingly
more voracious than adult males or juveniles. Many
species are cannibalistic.
2. Feeding Mechanisms
The feeding mechanisms of suspension-feeding
calanoids appear to be similar for marine and freshwater species. Food is brought in toward the copepod on
microcurrents created by rapid vibrations of the second
antennae, mandibular palps, first maxillae, and maxillipeds (Koehl and Strickler, 1981; Vanderploeg and Paffenhöfer, 1985). Smaller food particles are captured
passively and are funneled into the mouth through the
setae of the second maxillae (Vanderploeg and Paffenhöfer, 1985; Price and Paffenhöfer, 1986). During passive captures, the vibrations of the feeding appendages
continue uninterrupted. Larger food particles are captured actively with an outward fling of the second maxillae followed by an inward squeeze to remove the excess water surrounding the food particle.
When microzooplankton such as rotifers and nauplii are brought in on the feeding currents of suspension-feeding diaptomids, the copepods will often attack
these small animal prey from a distance with an active
thrust response in which the antennae and swimming
legs are used to orient and pounce toward the prey
(Williamson and Butler, 1986; Williamson, 1987;
Williamson and Vanderploeg, 1988). Particles less than
5 m are generally captured passively by diaptomids,
while those greater than 50 m are generally captured
actively or by attack; in intermediate size ranges, the
frequency of active captures increases with increasing
22. Copepoda
particle size (Vanderploeg and Paffenhöfer, 1985;
Williamson and Vanderploeg, 1988).
Many of the larger calanoid species such as Heterocope, Epischura, Limnocalanus, and some of the
large species of diaptomids have predatory tendencies
and feed on other zooplankton as well as algae. These
predators cruise through the water, attack their prey
with a pounce, and grasp them with their first and second maxillae. If a capture attempt fails, the copepod
may swim in a vertical loop to try again to capture the
prey (Kerfoot, 1978).
Cyclopoid copepods do not create currents to aid
their feeding but instead grasp their food directly with
their first maxillae or, to a lesser extent, with their second maxillae and maxillipeds. These appendages push
food between the mandibles. By oscillating rapidly during feeding bouts, the mandibles tear prey into pieces
and stuff them into the esophagus (Fryer, 1957a).
Cyclopoids detect their prey with the help of
mechanoreceptors on their first antennae. Prey which
generate much disturbance as they swim can be detected
at distances of several millimeters. Cyclopoids actively
orient and attack prey with considerable precision (Kerfoot, 1978). Larger prey such as cladocerans, copepods,
and chironomids may be handled for 30 min or more before ingestion (Fryer, 1957a; Gilbert and Williamson,
1978; Williamson, 1980). Small, generally less active
prey such as algae, protozoans, rotifers, and nauplii may
be detected only after contact, and are generally eaten
whole. Cyclopoid nauplii use their mandibles and second
antennae to capture and ingest food (Monakov, 1976).
Harpacticoid copepods are primarily surface-feeders that use their mouthparts for scraping food from a
diversity of substrates. Suspension-feeding has been reported in two primitive marine families, the Longipediidae and the Canuellidae (Hicks and Coull, 1983). In
surface-feeding, the second maxillae and second antennae grasp the prey while the maxillipeds aid in anchoring the copepod to the substrate. Once the food is
grasped, the first maxillae shred and stuff the food into
the vibrating mandibles. The second maxillae and maxillipeds are not used in food handling. Harpacticoid
nauplii have modified second antennae that are employed to grasp, masticate, and push food into the
mouth (Fahrenbach, 1962). The mandibles of nauplii
serve only for locomotion and not for feeding.
Suspension-feeding harpacticoids feed by lying on
their dorsal or lateral sides and vibrating their first and
second maxillae and mandibles. These vibrations create
feeding currents that bring water and food in toward
the mouth and out posteriorly along the ventral surface
of the copepod. Some harpacticoids can use both surface-feeding and suspension-feeding mechanisms
(Hicks and Coull, 1983).
929
3. Feeding Rates
Feeding rates are usually expressed quantitatively
as either clearance rates (expressed as volume of water
cleared of food per unit time) or ingestion rates (food
items ingested per copepod per unit time). Ingestion
rates generally depend upon prey density below a saturation food density referred to as the incipient limiting
concentration; above this density there is no further increase in ingestion rate. Clearance rates are density-independent below this saturation food concentration
and decrease above it. Feeding rates are calculated
based on the exponential equations first applied to zooplankton feeding by Gauld (1951). The standard experimental design is to fill a number of experimental
vessels with either food alone (controls) or food and
copepods (experimentals) and then incubate them for a
period of time on a rotating wheel to prevent settling
and minimize patchiness of the food and copepods.
Feeding rates can then be calculated as follows:
Ingestion rate (I) 5
Clearance rate (F)
Dct 2 Det
TN
(1)
V(ln Dct ln Det)
TN
(2)
where Dct and Det are the food densities in the control
and experimental vessels at the end of the incubation,
respectively, T the duration of the experiment, and N
the number of copepods per experimental vessel. The
average food density to which the copepods are exposed during the experiment is then:
d
Det Deo
V(ln Det ln Deo)
(3)
where d is the average food density, Det and Deo are the
final and initial food densities in the experimental vessels, respectively, V the volume of the experimental vessel, and I Fd.
An alternative method for estimating feeding rates
is to add different numbers of predators to the experimental vessels and use regression analysis to determine
clearance rates (Lehman, 1980; Landry and Hassett,
1982). This method assumes that changes in the experimental vessels are proportional to the number of
predators per vessel and to the incubation time according to the following equation:
ln Det Dct FNT
(4)
This relationship is derived from the conventional population growth rate equation (see Eq. 5). If Det /T is
plotted against N, the slope of the regression relationship is the clearance rate (F), the y intercept is an estimate of ln Deo, and standard regression statistics can be
used to determine the statistical significance of the
930
C. E. Williamson and J. W. Reid
regression relationship as well as the percentage of the
variation in prey densities that can be attributed to predation (or, how good was the experiment?). An assumption of both of the above methods of estimating
feeding rates is that Dct Deo , or, in other words, that
prey birth rates as well as nonpredatory death rates
were minimal.
Gut content analyses can also be helpful in distinguishing the diet of copepods. Copepods can be
squashed between a coverslip and a microscope slide
for a crude assessment of gut contents. A more quantitative analysis can be obtained by dissection. A pair of
fine minuten needles (insect pins) secured to the tips of
applicator sticks or glass pipets can be used to tease the
metasomal gut tube out of preserved copepods. The gut
is placed in a very small drop of water, covered with a
coverslip, and examined under a compound microscope before and after the addition of 5% sodium
hypochlorite (commercial bleach) (Williamson, 1984).
Clearance rates of predatory calanoids are generally in the range of 80 to 180, but values have been reported as high as 630 mL per predator per day, for
Hesperodiaptomus shoshone feeding on calanoid nauplii at 20°C (Maly, 1976) and over 1,000 mL per
predator per day, for Heterocope septentrionalis feeding on Daphnia pulex at 15°C (Luecke and O’Brien,
1983). Ingestion rates averaging as high as 18.2 prey
per predator per day have been reported for Hesperodiaptomus shoshone feeding on various crustacean prey
(Anderson, 1970); while rates of up to 54 prey per
predator per day have been observed for Heterocope
septentrionalis feeding on Bosmina longirostris at 15°C
(O’Brien and Schmidt, 1979). The benthic harpacticoid
Attheyella reduced bacterial densities in an Appalachian headwater stream by up to 58%; the greater
reductions were observed under lower flow conditions
(Perlmutter and Meyer, 1991).
The feeding rates of suspension-feeding calanoids
vary widely and probably depend largely on dietary
preferences and available food. Many investigators
have obtained clearance rates of only a few milliliters
per copepod per day or less for certain algae, and
calanoids are thus often thought to have clearance rates
that are much lower than similarly sized planktonic
cladocerans (Wetzel, 1983). However, when offered
high quality algae as food, diaptomids filter 19 – 25
milliliters per copepod per day, and weight-specific
rates are comparable to or greater than those of cladocerans (Bogdan and Gilbert, 1984; Vanderploeg et al.,
1984; Williamson and Butler, 1986). Although most of
the smaller diaptomids have historically been considered to be herbivorous, clearance and ingestion rates of
diaptomids on rotifers may be five to six times greater
than those for algae (Williamson and Butler, 1986).
The presence of algae reduces the predation rate of suspension-feeding diaptomids (Williamson and Butler,
1986), but does not similarly affect more predatory
species such as Epischura (Wong, 1981). In general, diaptomids feed more selectively than Daphnia (Richman
and Dodson, 1983; DeMott, 1988, 1989; Vanderploeg,
1994).
Cyclopoid ingestion rates are density-dependent
and may reach 42 prey per predator per day at prey
densities approaching 1000 prey / L (Williamson, 1984,
1986). Clearance rates may exceed 150 mL per predator per day (Williamson, 1983b). Ingestion rates, clearance rates, and prey selectivities may be influenced by
hunger levels of the predator, water temperature, and
the size, morphology, behavior, and density of the prey
(Williamson, 1980, 1986; Stemberger, 1986; Roche,
1987).
F. Population Regulation
1. Factors Controlling Population Density
Estimates of the per capita growth rates (r) of
copepod populations vary from less than 0.1 per day to
as high as 0.4 per day (Allan, 1976; Hicks and Coull,
1983). Doubling times (ln 2 / r) are generally on the
order of 1 to 2 weeks but may be less than 2 days under optimal conditions. These high reproductive rates
generate pronounced density oscillations in nature.
These oscillations, in conjunction with the 12 distinct
life-history stages, a small body size, and short generation times, make copepods good subjects for population studies.
Several hypotheses have been proposed to explain
the observed oscillations in copepod populations. The
most compelling of these include food limitation, temperature, and predation. Mate limitation may also be
important for diaptomids that must mate before the
production of each clutch (Williamson and Butler,
1987). The relative contributions of each of these factors vary between systems and are difficult to separate
in nature.
The physiological importance of temperature in
controlling rates of growth and development has been
demonstrated in numerous copepod species. Gamete
maturation, egg development, clutch production, and
instar development rates are all temperature-dependent
(Watras, 1983b; Hicks and Coull, 1983). In addition,
temperature may determine the seasonal timing of reproductive activity. A recent review of temperature responses of freshwater zooplankton indicates that many
species of copepods are cold-water species potentially
sensitive to climate warming (Moore et al., 1996).
Within the temperature range that permits reproductive
22. Copepoda
activity, food availability may limit the rates of reproduction and development of copepods by prolonging
interclutch duration and reducing clutch size (Woodward and White, 1981; Elmore, 1982, 1983). Response
to food limitation may be quite rapid in some species
(Williamson, et al., 1985).
The differential response of the reproductive
phases of copepods to food density, mate density, and
temperature may permit the analytical separation of
food and mate limitation from temperature effects in
some diaptomid copepods (Williamson and Butler,
1987). The transitions between reproductive phases are
largely regulated by food and mate availability and are
independent of temperature; hence, the proportion of
adult females in the different phases will reflect which
factors are limiting (Fig. 6). Food limitation can also be
examined with an index that quantifies both lipid storage and reproductive condition (Vanderploeg et al.,
1992b). This latter index is similar to that developed
for marine copepods (Marshall and Orr, 1952) and
freshwater cladocerans (Tessier and Goulden, 1982).
Optical – digital methods have also been used to quantify seasonal changes in lipid reserves in diaptomids
(Arts and Evans, 1991). They are much simpler than,
and compare favorably with, more expensive chemical
methods of analysis. The response of copepods to food
and mate limitation will vary with species and environmental conditions, and all of these indices must therefore be applied only with an adequate understanding of
the reproductive physiology of each species (Jersabek
and Schabetsberger, 1995).
In many populations the nauplii are the “bottleneck” stage that exhibits the highest mortality rate in
nature (Confer and Cooley, 1977; Feller, 1980). The
small size of the nauplii relative to copepodids may result in an increased vulnerability to both starvation
(due to higher weight-specific metabolic rates) and invertebrate predation, and consequently higher mortality rates for the nauplii (Williamson et al., 1985).
The relative abundance of calanoid versus cyclopoid copepods may change with the trophic status
of an ecosystem. Several investigators found that as
system productivity rose, the abundance of cyclopoid
copepods and cladocerans increased while calanoid
abundance either decreased or varied independently
(Byron et al., 1984; Pace, 1986; Stemberger and Lazorchak, 1994). Feeding and respiration rate experiments
with Daphnia and diaptomids indicated that food
quality may also play an important role in the relative
abundance of calanoids versus cladocerans (Richman
and Dodson, 1983; Lampert and Muck, 1985). Richman and Dodson (1983) suggested that calanoids will
predominate when either food quality is low or when
food density is either extremely high or extremely low.
931
Lampert and Muck (1985) suggested that diaptomids
are better adapted to low, fluctuating food densities
than is Daphnia, because the former allocates more energy to storage for survival while the latter puts its resources into reproduction under these food-limiting
conditions. There is some evidence that the threshold
food level for survival is lower for calanoid than for cyclopoid nauplii, and that this “bottleneck” stage may
be important in regulating the relative abundance of
these two taxa (Santer, 1994). Enclosure experiments in
450 L tanks indicate that the interactions between cyclopoids and calanoids are complex and may involve
predation on calanoids by cyclopoids, and depression
of food resources for cyclopoid nauplii by calanoids
(Soto and Hurlbert, 1991).
Predation may also influence copepod population
densities. Cannibalism is common among the more carnivorous species (Gabriel, 1985), and both vertebrate
and invertebrate predators prey on copepods. The inconsistent relationship between calanoid abundance
and lake trophic status discussed above may be due to
invertebrate predation on diaptomids (Edmondson,
1985), or possibly predation by the more carnivorous
cyclopoids on calanoids (Soto and Hurlbert, 1991;
Adrian, 1997). Other invertebrate predators, including
predatory copepods and larvae of the phantom midge
Chaoborus, may inflict substantial mortality on copepod populations (Luecke and Litt, 1987). The predatory cladoceran Bythotrephes cederstroemi was introduced into the Laurentian Great Lakes from Europe in
the 1980s, and has spread to nearby smaller lakes as
well. This exotic species preys on both adult copepods
and nauplii; and, while copepods seem to be some of
the less-preferred prey (Vanderploeg et al., 1993), some
cyclopoids such as Mesocyclops edax and Tropocyclops prasinus mexicanus have either declined or disappeared in lakes when Bythotrephes invades (Yan and
Pawson, 1997). Vertebrate predators, including fish,
birds, and larval and adult salamanders, may influence
the distribution and abundance of copepods in lakes
(Dodson and Egger, 1980; Zaret, 1980; Carter and
Goudie, 1986). Larger species such as Hesperodiaptomus may be excluded from lakes when fish are introduced (Donald et al., 1994). Vertebrate predators may
also be instrumental in the induction of diapause (Hairston and Walton, 1986) or diel vertical migrations
(Williamson and Magnien, 1982), and both vertebrate
and invertebrate predators may contribute to skewed
sex ratios in copepod populations on which they prey
(Maly, 1970; Hairston et al., 1983; Blais and Maly,
1993).
Heavy organic and nutrient loading and associated
reductions in dissolved oxygen may differentially reduce populations of different species of benthic
932
C. E. Williamson and J. W. Reid
harpacticoids, as well as reduce the abundance of
harpacticoids relative to other meiobenthos (Särkkä,
1992).
2. Methods for Analysis of Populations
The instantaneous per capita birth rates, death
rates, and growth rates of copepods can be estimated
using the egg ratio technique developed by Edmondson
(1960) and Edmondson et al. (1962), as modified by
Paloheimo (1974). The growth rate of the population
(r) is estimated from the population densities of adult
females at time zero (N0) and time t (Nt) with the following equation:
r
ln Nt ln N0
t
(5)
The birth rate (b) can be estimated from the egg ratio
(E) and the development time (D) from egg to adult as:
b
ln (E 1)
D
(6)
where E is the number of female eggs in the population
(assume a 1:1 sex ratio) divided by the number of adult
females in the population. The death rate (d) can then
be estimated from:
dbr
(7)
These estimates of r are for the midpoint between
the two sampling dates while the estimates of b are for
the given sample date. Therefore, when estimating d
from Eq. 6, the values of r should be linearly interpolated for the given date (Wyngaard, 1983). These methods assume a constant age structure and continuous
egg laying in the populations. Estimates of r with this
egg-ratio method are best applied to populations with
shorter developmental stages such as those found at
higher temperatures. An alternative approach which requires much greater resolution of the developmental
stage durations is cohort analysis (Hairston and
Twombly, 1985).
A careful analysis of the allocation of effort between and within sampling stations as well as between
subsamples is critical in order to maximize the accuracy of population estimates (Nie and Vijverberg,
1985). The variance associated with estimates of per
capita birth, death, and growth rates can be obtained
with either jackknife or bootstrap techniques (Meyer
et al., 1986).
G. Functional Role in the Ecosystem
Copepods make up a major portion of the biomass
and productivity of freshwater systems. In Mirror
Lake, New Hampshire, copepods contributed about
55% of the total zooplankton biomass over a three
year period (Makarewicz and Likens, 1979). Cyclopoid
copepods may contribute up to 77% and calanoid
copepods up to 49% of the macrozooplankton biomass
in some lakes (Pace, 1986). Benthic samples in a
second-order stream in North Carolina revealed biomass to be on the order of 22 mg dry mass/ m2, equivalent to densities of 3000 – 18,000 individuals / m2, and
cyclopoid densities of about 100 – 1600 individuals / m2
(O’Doherty, 1985, 1988). In the benthic communities
of lakes, it is not uncommon to find densities of copepods in excess of 10,000 and, occasionally, up to even
70,000 individuals / m2, with a biomass of 50 – 70 mg
dry weight individuals / m2 (Strayer, 1985).
In addition to their numerical contribution, copepods occupy an important intermediate position in
aquatic food chains (Kerfoot and DeMott, 1984). Their
generally omnivorous habits make them important in
energy transfer up the food chain; they also have the
potential to regulate the structure of phytoplankton
and zooplankton prey assemblages. By packaging bacterioplankton, phytoplankton, and microzooplankton
prey into larger particles (their own bodies) copepods
may actually increase food-chain efficiency in spite of
the energy lost by the additional intermediate trophic
level (Confer and Blades, 1975). Copepods may also
contribute to phosphorus regeneration in lakes (Bowers, 1986).
As predators, copepods may have a substantial impact on their prey populations. Predatory calanoids
and cyclopoids are intense, selective predators that can
alter the distribution and morphology of their prey.
Mesocyclops edax, an important cyclopoid predator in
the plankton, may crop up to 24% of its prey populations per day (Brandl and Fernando, 1979; Williamson,
1984). Populations of even some of the smaller suspension-feeding diaptomids have the potential to clear the
rotifers from a volume of water equivalent to the entire
volume of the lake in a single day (Williamson and Butler, 1986). Cyclops vicinus can prey on the rotifer Synchaeta at rates of up to 37 Synchaeta per day, rates that
suggest that this cyclopoid may be responsible for declines in this rotifer in Lake Constance in the spring
(Plassmann et al., 1997). Cyclopoid copepods prey selectively on, and may selectively inhibit the growth and
reproductive rates of, smaller cladoceran species as well
as juveniles within species (Gliwicz, 1994; Gliwicz and
Umana, 1994; Santer, 1993). In contrast, only very
large Daphnia appear to be susceptible to brood predation by copepods. Eudiaptomus gracilis and Acanthocyclops robustus copepodids have both been observed
to enter the brood cavities of Daphnia over 2.25 mm
long, where they prey on the eggs (Gliwicz and
22. Copepoda
Lampert, 1994; Gliwicz and Stibor, 1993). Some of the
large species of Hesperodiaptomus may regulate the
relative abundance of different rotifer species and the
clonal composition of Daphnia in arctic and alpine systems (Paul et al., 1995; Paul and Schindler, 1994; Wilson and Hebert, 1993). Several calanoid species in the
Great Lakes can feed on the larvae of the zebra mussel,
Dreissena polymorpha (Liebig and Vanderploeg,
1995).
Although smaller prey are generally preferred by
predatory copepods, the coincident vertical migration
of the copepods with some of the larger cladocerans
may cause ingestion rates to be disproportionally
greater on these otherwise undesirable prey types
(Williamson et al., 1989). Many prey species exhibit
elaborate morphological and behavioral defense mechanisms that suggest a strong coevolutionary relationship between copepod predators and their prey. The
hard loricas, spines, gelatinous sheaths, and rapid
escape responses of many prey species are all effective
at reducing predation by copepods (Williamson,
1983a, 1986, 1987; Stemberger, 1985; Roche, 1987).
Some of the morphological defenses are nongenetic
polymorphisms that are induced by chemical substances released by the copepods (Stemberger and
Gilbert, 1984).
Copepods are themselves important prey for other
copepods (Peacock and Smyly, 1983), Chaoborus larvae (Peacock, 1982; Wyngaard, 1983), and fish. Adult
copepods are preyed on by many planktivorous and
benthic-feeding fish, and the nauplii are particularly
important prey for larval fish (Guma’a, 1978; Hicks
and Coull, 1983).
Some of the larger cyclopoid copepods such as
Mesocyclops and Macrocyclops may serve as important
agents of biological control of pest insects such as mosquitos (Marten et al., 1994; Vu et al., 1998). Predation
rates on early instar mosquito larvae may be quite high
(Brown et al., 1991; Reid, 1989b), and cyclopoids can
be cultivated in large quantities for this purpose
(Marten et al., 1994; Suárez et al., 1992).
Cyclopoids and calanoids are also important intermediate hosts for many parasites, including flukes, nematodes, and tapeworms of fish, amphibians, birds,
and mammals. Some cyclopoids such as Mesocyclops
and Thermocyclops may serve as intermediate hosts of
the human parasite Dracunculus medinensis (guinea
worm) in the tropics. Ingestion of the larvae of this nematode in drinking water can cause serious illness or
death in humans in areas of equatorial Africa and India
(Steib and Mayer, 1988). The bodies of planktonic and
epibenthic copepods provide a substrate for the Vibrio
cholerae bacterium that carries cholera (Huq et al.,
1983).
933
Recent studies on the stoichiometric relationships
in planktonic populations have revealed that nutrient
ratios in copepods differ from other crustacean plankton. Since high growth rates require high ribosomal
RNA, high specific growth rates should be tied to a
higher percentage of phosphorus in zooplankton (Main
et al., 1997). Copepods tend to have a lower percentage of phosphorus in their bodies than many other
zooplankters (Sterner and Hessen, 1994). For example,
the mean atomic ratios of C:N:P are on the order of
212:39:1 for Acanthodiaptomus and 85:14:1 for
Daphnia (Andersen and Hessen, 1991). The implications are that these nutrient ratios will give diaptomids
a competitive advantage over cladocerans such as
Daphnia during periods of phosphorus limitation, and
also influence the importance of copepods in nutrient
cycling as well as a variety of other ecosystem processes
(Sterner and Hessen, 1994). Although seasonal changes
in zooplankton communities may contribute to temporal variations in levels of toxic trace metals such as cadmium, concentrations are much lower in copepods
than in cladoceran zooplankton (Yan et al., 1990).
IV. CURRENT AND FUTURE
RESEARCH PROBLEMS
One of the most striking things about freshwater
copepods is how little is known about them in comparison with either cladocerans or the marine copepods.
Basic information on things as elementary as morphological characterization, life cycles, reproductive biology, feeding ecology, physiology, and genetics is lacking
for most species. This is particularly true for the
harpacticoids. Outside of Europe, taxonomic knowledge is inadequate for many species, especially for the
non-planktonic species. Information of this sort will increase our understanding of copepods as vectors of diseases such as cholera and dracunculiasis, as well as
help us to realize their full potential in biological control of pest insects such as mosquitos (discussed above).
In addition to expanding our knowledge of copepods themselves, research on copepods can contribute
to our knowledge of more general ecological questions.
The small size, amenability to culture, discrete instars,
existence of diapause stages, and variability in lifehistory patterns permit many interesting questions in
population biology to be addressed. The intense and
selective feeding of copepods, as well as their importance as food for higher trophic levels, open the way
for studies in foraging behavior, community ecology,
and systems ecology.
While we are aware that food may limit copepod
populations, we know very little about the relative
934
C. E. Williamson and J. W. Reid
importance of food in controlling the presence or abundance of different copepod species in nature. We are
particularly naive about the interactive effects of environmental variables such as temperature, pH, solar ultraviolet radiation, the presence of alternate food, the
hunger level of the predator, predation risk, and various forms of pollution and environmental manipulation on copepods. With ecosystem disturbance being
the norm rather than the exception now across the
globe (Vitousek, 1994), it is critical that we gain a better understanding of how many of these disturbance-related abiotic and biotic variables affect key organisms
such as copepods.
Another interesting area of research to pursue concerns the relative abundance of cyclopoid versus
calanoid copepods across lakes of different trophic
status (discussed in Section III. F.1). The densities of cyclopoid copepods and most other zooplankton groups
(ciliates, rotifers, cladocerans) are positively correlated
with the primary productivity of lakes. This does not,
however, seem to be universally true for calanoid
copepods (Pace, 1986). What are the characteristics of
calanoids that permit them to respond differently
from most other groups to variations in primary
productivity?
Ecological genetics is another area that is advancing rapidly and is ripe for future study. The presence or
absence and timing of chromatin diminution varies
among species and may be useful in deciphering phylogenetic and evolutionary relationships among taxa
(Einsle, 1996; Leech and Wyngaard, 1996; Dorward
and Wyngaard, 1997). Little is known about the endogenous versus exogenous control of the wide range
of variations in life-history characteristics observed for
copepods in different systems or through different seasons. Many interesting evolutionary questions can be
asked about the timing and control of life-history
events of copepods due to the genetic isolation of populations and variability in selective pressures acting on
these populations among lakes. Of particular interest is
the recent finding that resting eggs may contribute substantially to the evolution and persistence of copepod
populations over long periods of time (Hairston et al.,
1995; Hairston, 1996), thus potentially permitting
copepods to survive bottleneck periods related to both
anthropogenic and natural disturbances.
Recent finds in North America of groundwaterrelated copepod species that are closely related to
Eurasian forms (e.g., Reid, 1998; Reid et al., 1999) are
throwing new light on biogeographical relationships
and the nature of evolution in copepods. While some
groups seem to be extremely conservative, apparently
having diverged little since the pre-Tethyan continental
separation, other species evolved more rapidly to take
advantage of new niches such as the cenotes of the
Yucatan Peninsula (Fiers et al., 1996).
V. COLLECTING AND REARING TECHNIQUES
A. Collection and Preservation
Planktonic copepods can be collected with either
integrating samplers, such as a plankton net, or tube
samplers (Lewis and Saunders, 1979), or with a variety
of point samplers such as Schindler traps and Van
Dorn bottles (Schindler, 1969). Sampling devices with
narrow openings and pump samplers may elicit avoidance responses in copepods and should be tested for
sampling efficiency before use. Quantitative samples
must be taken at night for species that exhibit nocturnal vertical migrations into the sediments. Benthic
copepods and the resting stages of planktonic species
can be collected with either Ekman grab or core samplers. Separation of the smaller benthic copepods from
the sediments presents difficulties. Sieving and sorting
are tedious and not very efficient for copepods. Density
gradient centrifugation seems to be a more promising
method (Strayer, 1985).
Copepods can be preserved in a 10% solution of
formalin (3.7% formaldehyde) or 70% ethanol. Formalin is less expensive but more toxic, and tends to
cause specimens to get brittle over time due to the low
pH. Buffering formalin with a saturated solution of borax (sodium tetraborate, Na2B4.1H2O) will improve
preservation quality by making the specimens less brittle, but the high pH may also lead to more rapid color
loss. A 70% solution of ethanol is actually a better
preservative but is also more expensive, bulky, and may
create difficult eddies in open counting chambers, making quantification extremely difficult. Prior narcotization of copepods in carbonated water may enhance the
retention of the gut contents of some species when they
are placed in formalin (Gannon and Gannon, 1975)
but is not always necessary (Williamson, 1984). The retention of egg clutches may be enhanced by prior narcotization in a 1:10 solution of chloroform: ethanol
(Williamson and Butler, 1987). The chilling and addition of 40 g of sucrose / L of formalin commonly employed in the preservation of cladocerans (Prepas,
1978) is not required, but it also has no adverse effect
on copepods. The adult stages of copepods can be
counted in a Bogorov chamber under a dissecting microscope (Gannon, 1971), while the nauplii are more
easily counted in a Sedgwick – Rafter chamber under a
compound microscope.
Several techniques have been developed for the separation of copepods in samples. Many live copepods can
be separated from cladocerans with siphon tubes that
22. Copepoda
remove the slower cladocerans but not the evasive copepods; or, one can take advantage of the greater resistance of copepods to pH shock (Bulkowski et al., 1985).
Live copepods can be separated from dead copepods by staining with neutral red (Dressel et al., 1972;
Flemming and Coughlan, 1978). A stock solution is
made with 0.5 g of neutral red powder per liter of distilled water. This stock solution is then added to the
sample containing the copepods in a 2:100 ratio for a
staining period of up to 1 h before preservation in formalin. The prior addition of 4 g of NaOH per liter of
formalin reduces the leaching of the stain out of the
copepods for up to 28 weeks. The high pH makes the
stain much less visible, however, and glacial acetic acid
must be used to titrate the sample back down to an
acid pH to regenerate the deep red color before sorting.
Techniques have also been developed for the critical examination of the anatomy of copepods. The external anatomy, including sensilla and other delicate integumental organs, can be prepared by critical point
drying and plating for examination under a scanning
electron microscope, or cleared, stained, and examined
under a light microscope (Fleminger, 1973; Friedman
and Strickler, 1975). The gonads of some copepods can
be selectively stained with fast green for easier visualization (Batchelder, 1986), and techniques are available
for the histopathological examination of copepods
(Barszcz and Yevich, 1976).
935
Freshwater diaptomids ranging from the smaller
Leptodiaptomus minutus to the larger more omnivorous Hesperodiaptomus shoshone can be raised
through multiple generations on a diet of Cryptomonas
alone. Cyclopoid copepods are generally best reared on
a combination of plant and animal foods. The nauplii
will survive and grow well on a variety of algae including Cryptomonas and Scenedesmus, while animal food
is usually necessary for the growth and survival of late
copepodids and adults and for successful reproduction
of some species (Jamieson, 1980b; Wyngaard and
Chinnappa, 1982; Hansen and Santer, 1995; Hart and
Santer, 1994). Algae as well as protozoa and invertebrates may still be an important food source for these
older predatory stages (Santer, 1996). In culture, the
dinoflagellate Ceratium furcoides supports the growth
and reproduction of several genera of cyclopoid copepods but not those of the calanoid Eudiaptomus
gracilis (Santer, 1996; Hopp et al., 1997). The early
copepodid stages may need a mixture of plant and animal foods. Artemia nauplii are the most common and
convenient source of animal food for cyclopoids, but
calanoid copepods or soft-bodied cladocerans and rotifers may also be used. Protozoans are a good food
source, but they may lead to bacterial contamination
unless care is taken to rinse and remove the organic
medium or use an inorganic medium for culture. Mass
rearing of cyclopoids is also possible (Suárez et al.,
1992).
B. Rearing Techniques
Inorganic medium, spring water, or filtered lake
water can be used in conjunction with a variety of
foods to culture copepods. The utmost care should be
taken to avoid glassware contaminated with soaps,
acids, formaldehyde, or other toxins. Certain plastic
containers may also be unsuitable. An easily rinsed detergent such as 7X®, should be used to clean all glassware thoroughly. [The source of 7X® is ICN Biomedicals, Inc., Costa Mesa, CA; phone:800-854-0530;
www.icnpharm.com.] Standard detergents may cause
high mortalities. Dilute acids may also be satisfactory
for washing glassware (Wyngaard and Chinnappa,
1982), but thorough soaking or extra rinsing is suggested. Water quality may also be enhanced by running
distilled water through a mixed-bed resin before use.
All glassware should be sterilized between uses to avoid
contaminant buildup.
Harpacticoids have been cultured on a wide range
of artificial and natural foods, including baker’s yeast,
dried fish food, algae, bacteria, and dried, mashed, or
rotting animals, eggs, and plants (Hicks and Coull,
1983). Multiple foods are often more successful than a
single food type.
VI. IDENTIFICATION TECHNIQUES
A. Dissection Techniques for Identification
Dissection is usually required for positive identification of copepods. Small insect pins (minuten nadeln,
available from entomological supply houses) can be
mounted on the end of wood applicator sticks or
melted glass pipets for use in dissection. The dissection
should be performed on a glass microscope slide so
that a coverslip can be placed over the final preparation
for examination under a compound microscope. The
dissection is most easily performed by placing the copepod on its side in a drop of medium, and placing one
pin in the dorsal part of the thorax to hold it down. A
second pin is then used to tease off first the urosome,
then each thoracic segment and its associated pair of
legs — one segment at a time, and finally the first antennae. The chance of confusing legs is lessened if the first
cut is made between the second and third legs, and the
posterior part of the animal transferred to a second
drop of medium on the same slide. Dissection and handling of the pieces is easier when performed in a drop
936
C. E. Williamson and J. W. Reid
of glycerin or other viscous, water-miscible mounting
medium such as Hoyer’s or CMC-9. Placing a copepod
directly in full strength medium may cause osmotic
stress and deformation. This problem can be avoided
by placing the copepod in a well slide in a dilute mixture of about one drop of the medium to 5 – 10 drops
of water and letting the water evaporate. Gentle warming will speed evaporation.
Semi-permanent mounts can be made by sealing a
glycerin coverslip with clear fingernail polish. One of
several standard mounting media can be used for more
permanent mounts. A small amount of neutral red dye
in glycerin or a mounting medium with a dye will color
the dissected parts so that they are easier to locate after
dissection. The dissection can be performed right in the
viscous medium, the dissected appendages appropriately arranged in order, and the coverslip put in place.
When using a mounting medium other than glycerin it
is useful to let the mounting medium dry somewhat after dissection before adding more medium and adding
a coverslip. This helps the dissected parts stay put. If
the medium is allowed to dry for a week and the edges
sealed with fingernail polish or spar varnish, the mount
will last indefinitely.
For more details on techniques for copepod preparation, dissection, staining, mounting, and preservation,
the reader is referred to an unpublished pamphlet prepared by J. W. Reid and available in the C. B. Wilson
Copepod Library at the National Museum of Natural
History, Smithsonian Institution, Washington, DC.
B. Characteristics for Distinguishing the Free-Living
Freshwater Orders
Copepods are most easily identified from adult
specimens, although information is available for the
separation of the immature stages of some species
(Comita and Tommerdahl, 1960; Czaika and Robertson, 1968; Katona, 1971; Comita and McNett, 1976;
Shih and Maclellan, 1977; Bourguet, 1986; PinelAlloul and Lamoureux, 1988a, b). Growth rates and
some external morphological characteristics of copepods can be analyzed by examining the highly transparent molted exoskeletons (exuviae) (Twombly and
Burns, 1996). In freshwater habitats, the three major
copepod orders can be distinguished by a number of
characteristics of the adults; the length of the first antennae and taper of the body are perhaps the most useful of these (Fig. 1). The first antennae of harpacticoids
are very short (5 – 9 segments in the female) and rarely
extend past the posterior end of the cephalothorax. In
cyclopoids the first antennae are intermediate in length
(6 – 17 segments in the female), and rarely extend beyond the posterior end of the prosome. In calanoids the
first antennae have 23 – 25 segments in the female and
generally extend well beyond the end of the prosome to
the urosome or even the caudal setae. The body of
harpacticoids generally shows only a slight taper from
the anterior to the posterior, so that the metasome and
urosome are of similar widths. In cyclopoids and
calanoids the metasome is substantially wider than the
urosome. In cyclopoids there is a strong but gradual taper from front to back; while in calanoids there is a
more abrupt change in body width where the metasome joins the urosome. The body shape of harpacticoids is quite diverse and varies greatly with their habitat. For example, interstitial species are smaller and
more elongate, burrowing species are larger and
broader, while epibenthic species are large and variable
in their body shape (Hicks and Coull, 1983). The distinguishing features of the gelyelloids (all three species
of which are under 0.5 mm in body length) are the absence of the fourth (in the European species) and fifth
swimming legs, and the fusion of the coxae of the first
three pairs of swimming legs; their general body shape
is narrow near the midsection (Fig. 1E).
Other factors that distinguish the four free-living
orders include general habitat preferences and the number of egg sacs. Calanoids are almost exclusively planktonic and either carry a single egg sac medially or release their eggs directly into the water. Cyclopoids are
primarily benthic, although several planktonic species
are widespread and abundant, and carry two egg sacs
attached laterally to their genital segment. Harpacticoids are generally smaller than either calanoids or cyclopoids, are generally benthic or associated with other
substrates or particles, and usually carry a single egg
sac medially. Wilson and Yeatman (1959) provided a
useful table that summarizes other differences among
these three orders. Smallest of all are the gelyelloids,
which occur exclusively in subterranean karst or
stream hyporheic sediments; the nature of their egg
sacs is unknown.
Adult copepods are sexually dimorphic and sexes
can be distinguished on the basis of their relative size,
as well as certain primary or secondary sexual characteristics. Females are generally larger than males, and
mature gonads, spermatophores, or externally-carried
egg sacs are often visible. The subitaneous and resting
eggs of diaptomids can be distinguished from each
other in preserved samples by the presence of a space
between the chorion and the vitelline membrane in the
subitaneous eggs, and its absence in resting eggs
(Lohner et al., 1990). Adult male copepods have one
(calanoids) or two (harpacticoids, cyclopoids, and
gelyelloids) geniculate first antennae that are modified
for grasping the females during mating (Fig. 1 and 3).
After preservation the geniculate antennae of males
22. Copepoda
often become sharply bent, or elbowed, so sexes are
quite easy to distinguish. In harpacticoids, males can be
separated from females by their first antennae, which
have fewer and broader segments, and often more
densely clumped setae. Other characteristics such as
morphological differences in the caudal rami and legs
2 – 4 can be used to distinguish male and female
harpacticoids in many genera. In addition, female
harpacticoids generally have a more highly developed
fifth leg, and the sixth leg tends to be tiny. Characteristics that distinguish sex are generally not visible in any
of the earlier larval stages and are less conspicuous in
stage C5.
Adult copepods can be distinguished from immature copepodids either by size, or by the primary or secondary sexual characteristics described above. In adult
cyclopoids and harpacticoids the last segment of the
urosome is shorter than, or similar in length to, the next
to last (penultimate) segment, while in immatures the
last urosomal segment has not yet divided and is about
twice as long as it is wide (Fig. 5). Characteristics of the
genital segment such as a ventral swelling or more angular side profile in the adults may also be helpful in
separating adult females from immatures. The larval
stages within each order can be differentiated on the basis of body shape and the number and structure of the
appendages. Several individuals of each sex and stage
should be compared to establish valid criteria for distinguishing the sexes and stages of each species examined.
C. Identification of the Continental
Free-Living Copepoda
1. Introduction
Here we provide keys to the orders and genera, followed by tables that list the families, genera, and
species of copepods recorded from continental and
nearshore oligohaline habitats in North America. Keys
and revisions for individual genera are cited in the tables, while some of the broader, more comprehensive
keys and taxonomic references are listed in the paragraphs below. The taxonomy and systematics of copepods are active areas of research. In North America,
the faunas of ephemeral, subterranean, interstitial, and
semiterrestrial habitats are still poorly known. Moreover, refinements of taxonomic methods have led to redefinition of many taxa (Reid, 1998). Investigators
should consult with appropriate specialists and archive
reference specimens.
Dussart and Defaye (1995) provided an introduction to the taxonomy and systematics of the continental Copepoda, with diagnoses and keys to most genera.
For North American free-living continental copepods,
937
the most complete and best-illustrated keys are those
by Wilson and Yeatman (1959). The keys by Pennak
(1989) include most species of Calanoida and some Cyclopoida. Basic general taxonomic references for marine harpacticoids include the works of Coull (1977),
Lang (1948, 1965), and Huys et al. (1996).
There are several useful keys and checklists for different regions of North America. Works treating
calanoids, cyclopoids, and harpacticoids include those
of Cole (1959) for Kentucky, Robertson and Gannon
(1981) for the Great Lakes, Clamp et al. (1999) for
North Carolina, and Chengalath and Shih (1994) for
northwestern North America. Lists for calanoids and
cyclopoids include those of Harris (1978) for northern
Mississippi, Reed (1963) for northern Canada, and
Smith and Fernando (1978) for Ontario. Cole (1961,
1963, 1966) summarized records of Diaptomidae
(Calanoida) from Arizona, and Robertson (1970,
1972, 1975) listed diaptomids from Oklahoma. Czaika
(1982) provided keys and tables for adults and developmental stages of planktonic calanoids and cyclopoids
of the Great Lakes. Regional works on Cyclopoida include the key by Torke (1976) for Wisconsin and the
checklists by Bunting (1973) for Tennessee and Reid
and Marten (1995) for Louisiana and Mississippi.
Hudson et al. (1998) provided a checklist of cyclopoids
and harpacticoids of the Great Lakes and a key for the
cyclopoids.
Checklists for Mexico were furnished by SuárezMorales and Reid (1998) and by Reid (1990) for Mexico, Central America, and the Antilles. Suárez-Morales
et al. (1996) provided keys and taxonomic discussion for
species of the Yucatan Peninsula, as well as a Spanishlanguage general introduction to the study of continental
copepods.
Two public resources for copepod studies are the
C. B. Wilson Copepod Library at the National Museum of Natural History, Smithsonian Institution,
Washington DC., and the Monoculus Library at Oldenburg University in Germany. The Wilson Library
contains some 10,000 works, indexed by author and
species. Its website (http: / /www.nmnh.si.edu / iz / copepod) contains five databases: (1) a bibliography of all
known literature on Copepoda and Branchiura; (2)
taxonomic lists of families, genera, and species; (3) a
world list of researchers; and (4) the taxonomic
“types” of Copepoda and Branchiura held in the National Museum of Natural History. Contact F. D. Ferrari, T. C. Walter, or J. W. Reid for more information.
The Monoculus Bibliography Project also holds about
10,000 works, many entered in its database; contact H.
K. Schminke, Fachbereich 7-Biologie, Universität Oldenburg, Postfach 2503, D-26111 Oldenburg, Germany
for details.
938
C. E. Williamson and J. W. Reid
The true continental Calanoida are dominated by
the families Centropagidae, Diaptomidae, and Temoridae, and the Cyclopoida by the family Cyclopidae. The
continental harpacticoid fauna consists mainly of the
families Canthocamptidae, Parastenocarididae, and
Phyllognathopodidae, but the primarily marine families
Ameiridae, Huntemanniidae, and Laophontidae also
contain some true freshwater species. Several additional calanoid, cyclopoid, and harpacticoid families
include euryhaline species that may wander into coastal
freshwaters, but these are not listed in the tables. Use-
ful taxonomic references for euryhaline species include
C. B. Wilson (1932) for calanoids and cyclopoids; Walter (1989) for Pseudodiaptomus; Rocha (1991), Rocha
and Hakenkamp (1993), Rocha et al. (1998), and M.
S. Wilson (1958) for Halicyclops; and Coull (1977),
Lang (1965), and Huys et al. (1996) for harpacticoids.
2. Key to Orders and Families of the Free-Living
Continental Copepoda of North America
[Note that there are 3 or 4 couplets in some parts
of this taxonomic key.]
1a.
Body constricted, or major body joint between somites bearing 4th and 5th legs; first antenna of 6 – 17 segments.........................2
1b.
Body constricted between somite bearing 5th leg and genital segment; first antenna long, of 23 – 25 segments (Figs. 1A, 2, 5)
Order Calanoida, ...............................................................................................................................................................................4
1c.
No distinct body articulation between prosome and urosome; first antenna with 14 poorly defined segments; swimming legs
without couplers (intercoxal sclerites), and joined at base (Fig. 1E) .........................................................................Order Gelyelloida
2a(1a).
Second antenna of 3 or 4 segments, not prehensile; first antenna of with 6 – 17 segments...............................................................3
2b.
Second antenna of 3 segments, with large prehensile terminal claw; first antenna of with 6 segments .............................................
Order Poecilostomatoida (Ergasilidae) .................................................................................................................................................
3a(2a).
Fifth leg small, of 1 – 3 short segments, or represented only by setae; segment 1 not enlarged on inner margin (Figs. 1B, 2, 5)
...........................................................................................................................................................................Order Cyclopoida 24
3b.
Fifth leg broad, of 1 – 2 segments, segment 1 (baseoendopod) enlarged on inner margin (Figs. 1C,D, 2, 3) ..........................................
Order Harpacticoida, .......................................................................................................................................................................44
ORDER CALANOIDA
4a(1b).
Caudal ramus with 3 or 5 well-developed setae (plus 1 or 2 shorter, slender setae) ....................................................................5
4b.
Caudal ramus with 4 well-developed setae (plus shorter, slender outer and inner setae); legs 1 – 4 endopods of 1, 2, 3, and 3
segments respectively; first antenna not geniculate; leg 5 lacking...................................................................Aetideidae Senecella
5a(4b).
Caudal ramus with 3 well-developed terminal setae (plus reduced or spiniform outer seta and slender, dorsally placed inner
seta) ...................................................................................................................................................................................................6
5b.
Caudal ramus with 5 well-developed setae (plus slender, dorsally placed inner seta) ..................................................................7
6a(5a).
Caudal ramus , outer seta slender, about as long as ramus; leg 5 and left leg 5 , last segment with long apical spine; urosome symmetrical ........................................................................................................................................Temoridae Heterocope
6b.
Caudal ramus , outer seta shorter than ramus, and or spiniform; leg 5 and left leg 5 , last segment without long apical
spine; urosome asymmetrical, with various processes on right side ................................................................Temoridae Epischura
7a(5b)
Caudal ramus long (more than 3 times longer than wide) ..........................................................................................................8
7b.
Caudal ramus short (3 times or less than 3 times longer than wide) .........................................................................................10
8a(7a).
Maxillipeds elongate (about 2 times body width) in lateral view; legs 1 – 4, endopods all of 3 segments ............................................9
8b.
Maxillipeds not elongate (subequal to body width) in lateral view; legs 1 – 4, endopods of 1, 2, 2, and 2 segments, respectively; leg 5
lacking endopods ....................................................................................................................................Temoridae Eurytemora
8c.
Maxillipeds reduced; legs 1 – 4, endopodites all of 2 segments; leg 5 lacking endopodite..............................................................
Acartiidae ....................................................................................................................................................................Acartia sinensis
8d.
Maxillipeds not elongate; legs 1 – 4, endopodites all of 3 segments............................................Pseudodiaptomidae Pseudodiaptomus
9a(8a).
Caudal ramus with tiny spines on surface; body somite bearing leg 5 with no spines; right leg 5 , exopod segment 2, coxa with,
basis without medial process; leg 5 , exopod segment 2 with outer spine, and endopod segment 1 with inner seta ...........................
Centropagidae ................................................................................................................................................................Limnocalanus
22. Copepoda
939
9b.
Caudal ramus with no surface spines; body somite bearing leg 5 with 1 spine on each posterior corner; right leg 5 , coxa without
medial process, and basis with medial process; leg 5 , exopod segment 2 with no outer spine, and endopod segment 1 with no inner seta ......................................................................................................................................................Centropagidae Sinocalanus
10a(7b).
Caudal ramus , lengths of setae unequal, fourth seta from outer margin longer and stouter than others; legs 1 – 4, endopods all
with 3 segments; leg 5 , endopods 3-segmented, last segments with 6 long setae ...........................................................................
Centropagidae ...............................................................................................................................................................Osphranticum
10b.
Caudal ramus , lengths of setae nearly equal; leg 1 endopod of 2 segments, legs 2 – 4 endopods of 3 segments; leg 5 , endopods 1- or 2-segmented, last segments with 0 – 2 short setae ..................................................................................Diaptomidae 11
11a(10b).
Last thoracic segment with rounded or spiniform dorsal process .................................................................................................12
11b.
Last thoracic segment of female with no dorsal process ..................................................................................................................13
12a(11a).
Right leg 5 , terminal claw of exopod segment 2 slender, at least 1.5 times longer than segment; leg 5 , endopod with row of fine
terminal hairs and 2 long terminal spines ................................................................................................................Mastigodiaptomus
12b.
Right leg 5 , terminal claw of exopod segment 2 slender, usually twice or more length of segment; leg 5 , endopod usually with
fine terminal hairs only..................................................................................................................................Arctodiaptomus (in part)
12c.
Right leg 5 , terminal claw of exopod segment 2 stout and about as long as segment; left leg 5 , terminal process strongly corrugated; leg 5 , endopod with row of terminal hairs and 1 short spine ..........................................................................Sinodiaptomus
13a(11b).
Right first antenna , terminal segment with end rounded, bearing only setae.................................................................................14
13b.
Right first antenna , terminal segment with small clawlike process on distal end .................................................Acanthodiaptomus
14a(13a).
Left leg 5 , distal process of exopod not distinct, i.e., exopod of 2 segments .................................................................................16
14b.
Left leg 5 , distal process of exopod distinct, i.e., exopod of 3 segments........................................................................................15
15a(14b).
Right first antenna , antepenultimate segment with short stout curved spinous process on distal end; leg 5 , exopod segment 3
absent (with 2 remnant spines inserted directly on segment 2).................................................................................Onychodiaptomus
15b.
Right first antenna , antepenultimate segment with no process; leg 5 , exopod segment 3 present, distinct..........Nordodiaptomus
16a(14a).
Right first antenna , antepenultimate segment with distal spinous process.....................................................................................18
16b.
Right first antenna , antepenultimate segment with at most narrow hyaline lamella......................................................................17
17a(16b).
Both first antennae and left first antenna , with 2 setae on segment 11 and 2 setae on segments 15, 16, and 17
....................................................................................................................................................................................Mixodiaptomus
17b.
Both first antennae and left first antenna , with 1 seta on each of segments 11 and 15 – 17 .................................Skistodiaptomus
18a(16a).
Right first antenna with simple, rodlike or dentiform process on antepenultimate segment ..........................................................19
18b.
Right first antenna with comb-shaped (corrugated) process on antepenultimate segment; leg 5 with segment 2 shorter than segment 1 and with 1 rounded, haired pad; leg 5 , endopod usually with terminal hairs only .........................Arctodiaptomus (in part)
19a(18a).
Right first antenna , antepenultimate segment with short dentiform process (usually shorter than penultimate segment) ..............21
19b.
Right first antenna , antepenultimate segment with long dentiform process (usually longer than penultimate segment) ................20
20a(19b).
Left leg 5 , segment 2 usually longer than segment 1, bearing two distinctly divided, more or less hairy pads; leg 5 , exopod segment 3 present, distinct ..........................................................................................................................................Hesperodiaptomus
20b.
Left leg 5 , segment 2 shorter than segment 1, bearing 2 indistinctly divided pads; leg 5 , exopod segment 3 absent, with only 2
remnant spines, inserted directly on segment ................................................................................................Leptodiaptomus (in part)
21a(19a).
Both first antennae and left first antenna , setae on segments 17, 19, 20, and 22 with normal tapered ends (not hooked); left leg
5 , exopod segment 2 with lateral seta short, not extending past end of segment...........................................................................22
21b.
Both first antennae and left first antenna , setae on segments 17, 19, 20, and 22 with ends stiffly hooked; left leg 5 , lateral
seta of exopod 2 very long .........................................................................................................................................Aglaodiaptomus
22a(21a).
Both first antennae and left first antenna , segment 11 with 1 seta.............................................................................................23
22b.
Both first antennae and left first antenna , segment 11 with 2 setae..............................................................................Diaptomus
23a(22a).
Left leg 5 , exopod segment 2 with both terminal process and inner spine short and stout; leg 5 , exopod segment 3 absent, with
only 2 remnant spines, inserted directly on exopod 2 ...................................................................................Leptodiaptomus (in part)
23b.
Left leg 5 , exopod segment 2 with both terminal process and inner spine relatively long and slender; leg 5 , exopod segment 3
present and distinct .........................................................................................................................................................Eudiaptomus
940
C. E. Williamson and J. W. Reid
ORDER CYCLOPOIDA
24a(3a).
Mandibular palp consisting of tiny segment bearing 1 – 3 setae, or mandibular palp entirely absent ..............................Cyclopidae 25
24b.
Mandibular palp large, complex, with many plumose setae .........................................................................Oithonidae Limnoithona
25a(24a).
Leg 5 completely fused to body somite and bearing 3 setae and or spines; first antenna of 9 – 11 (usually 11) segments ..............26
25b.
Leg 5 consisting of 1 or 2 distinct segments; first antenna of 6 – 17 segments ...............................................................................28
25c.
Leg 5 consisting of 3 distinct segments; first antenna of 16 segments ..........................................................................Orthocyclops
26a(25a).
Leg 5 reduced to narrow plate or knobs, bearing 3 tiny setae; legs 1 – 4, endopods and exopods of 2 or fewer segments .................27
26b.
Leg 5 consisting of broad plate, bearing 2 strong inner spines and 1 long outer seta; legs 1 – 4, endopods and exopods all of 3 segments .................................................................................................................................................................................Ectocyclops
27a(26a).
Anal operculum large, triangular, with dentate margin; leg 4 coxa, inner corner lacking seta; leg 3 , endopod terminal segment
with modified apical spine Bryocyclops ................................................................................................................................................
27b.
Anal operculum small, quadrate, with smooth margin; leg 4 coxa with seta on inner corner; leg 3 , endopod terminal segment
with apical spine simple Stolonicyclops ................................................................................................................................................
28a(25b).
Leg 5 consisting of 1 distinct, broad segment and bearing 1 inner spine and 2 outer setae ...............................................................29
28b.
Leg 5 consisting of 1 or 2 distinct, usually narrow segments (proximal segment may be fused to body somite, but remnant seta of
this segment always present), segment 2 with 1 – 3 spines and setae (if 3, there is 1 middle spine between 2 setae)...........................33
29a(28a).
First antenna of 8, 11, or 12 segments; small species (usually under 1.4 mm long).......................................................................30
29b.
First antenna of 17 segments; large robust species (1.7 – 2.9 mm long)........................................................................Homocyclops
30a(29a).
First antenna of 12 segments ........................................................................................................................................................31
30b.
First antenna of 8 or 11 segments ....................................................................................................................Paracyclops (in part)
31a(30a).
Caudal ramus , with tiny spines next to lateral and outermost terminal setae only........................................................................32
31b.
Caudal ramus with longitudinal row of tiny spines on outer margin; caudal ramus at least 4 times longer than wide
.............................................................................................................................................................................................Eucyclops
32a(31a).
First antenna , last 2 segments not more that 2 times longer than wide; caudal ramus about 2.5 times longer than wide
............................................................................................................................................................................Paracyclops (in part)
32b.
First antenna , last 2 segments more than 3 times longer than wide; caudal ramus about 3 times longer than wide
........................................................................................................................................................................................Tropocyclops
33a(28b).
Leg 5 with 2 distinct segments..........................................................................................................................................................34
33b.
Leg 5 with 1 distinct segment ...........................................................................................................................................................40
34a(32a).
Leg 5, distal segment narrow, bearing 1 – 2 spines or setae ...............................................................................................................35
34b.
Leg 5, distal segment broad, bearing 3 spines and setae ..................................................................................................Macrocyclops
35a(34a).
Distal segment of leg 5 bearing 1 apical seta and 1 long inner marginal or subapical spine (length of spine more than 4 times length
of segment); first antenna always of 17 segments .............................................................................................................................36
35b.
Distal segment of leg 5 bearing 1 apical seta and (usually) 1 rather short inner marginal or subapical spine (length of spine not more
than twice length of segment); first antenna of 17 or fewer segments ...............................................................................................37
36a(35a).
Leg 5, inner spine of distal segment inserted about midlength of inner margin of segment; first antenna, last segment usually 3 or
more times longer than wide ............................................................................................................................................Mesocyclops
36b.
Leg 5, inner spine of distal segment inserted subterminally on segment; first antenna, last segment about twice longer than wide
.....................................................................................................................................................................................Thermocyclops
37a(35b).
Leg 5, distal segment with apical seta and (usually) small slender subapical spine on inner margin; caudal ramus without longitudinal dorsal ridge, and inner surface with or without hairs .................................................................................................................38
37b.
Leg 5, distal segment with apical seta and large spine (about equal in length to distal segment) attached at middle of inner margin of
segment; caudal ramus usually with longitudinal dorsal ridge, and inner margin hairy; first antenna of 14 – 17 segments........Cyclops
38a(37a).
Leg 5, segment 2 with apical seta and small spine or spur on middle of inner surface, or somewhat longer subapical spine (usually
shorter than segment 2); caudal ramus with or without hairs on inner surface .................................................................................39
38b.
Leg 5, segment 2 with inner subapical spine longer than segment 2; caudal ramus usually without hairs on inner surface
...........................................................................................................................................................................................Diacyclops
22. Copepoda
941
39a(38a).
Leg 5, segment 2 with apical seta and small spine or spur (usually a spur, continuous with segment) at about middle of inner surface; caudal ramus, inner surface hairy; first antenna of 17 segments ...............................................................................Megacyclops
39b.
Leg 5, segment 2 with apical seta and small spine slightly distal to middle of segment, or almost apical; caudal ramus with or without hairs on inner margin; first antenna of 11 – 17 segments ........................................................................................Acanthocyclops
40a(37b).
Leg 5, free segment narrow and cylindrical, or only slightly longer than wide, with inner apical spine or seta (if present) inserted
near outer apical seta .......................................................................................................................................................................41
40b.
Leg 5, free segment very broad, with inner apical spine inserted far from outer apical seta ................................................Apocyclops
41a(40a).
Leg 5, free segment narrow, cylindrical, with inner spine (if present) tiny and inserted subterminally or on inner surface of segment;
proximal segment completely absent, except for remnant seta..........................................................................................................42
41b.
Leg 5, free segment only slightly longer than wide, with slender inner terminal seta inserted next to longer outer terminal seta; proximal segment obvious although continuous with body somite ..........................................................................................................43
41c.
Leg 5, free segment only slightly longer than wide, with stout inner apical spine and outer apical seta both inserted terminally; proximal segment completely absent, except for remnant seta..................................................................................................Metacyclops
42a(41a).
Legs 1 – 4, couplers and coxa-basis very wide (about 2.5 times wider than long); legs 2 – 4, basis with rows of tiny spines on inner
expansion; leg 4 endopod, outer terminal spine tiny, less than 1 /6 length of inner terminal spine ..................................Cryptocyclops
42b.
Legs 1 – 4, couplers and coxa-basis less wide (about 2 times wider than long); legs 2 – 4, inner expansion of basis with fine hairs; leg
4 endopod, outer terminal spine at least 1 / 2 length of inner terminal spine ...................................................................Microcyclops
43a(41b).
Proximal segment of leg 5 present as tiny knob set close to free segment; legs 1 – 4 with 3- or 2-segmented rami; if exopodites of
2,2,3,3 segments, then exopodite segment 1 with medial (inner) seta................................................................................Rheocyclops
43b.
Proximal segment of leg 5 present as large knob set apart from free segment; legs 1 – 4 with endopodites 2-segmented and exopodites of 2,2,3,3 segments; exopodite segment 1 without medial (inner) seta ...................................................................Itocyclops
ORDER HARPACTICOIDA
44a(3b).
Somite bearing leg 1 completely fused with cephalothorax; maxilliped prehensile (ending in long claw), never flat and leaflike
.........................................................................................................................................................................................................45
44b.
Somite bearing leg 1 not fused to cephalothorax; maxilliped flat and leaflike, with several setae..........................................................
Phyllognathopodidae Phyllognathopus ................................................................................................................................................
45a(44a).
Leg 1 endopod prehensile (with dist almost segment long) or not, with 2 or more terminal setae (which may be stout)...................46
45b.
Leg 1 endopod prehensile, much longer than exopod, ending in 1 stout claw ......................................Laophontidae Onychocamptus
46a(43a).
Leg 1 exopod of 3 segments, middle segment with no spine on outer margin; body vermiform (narrow, cylindrical) .......................47
46b.
Leg 1 exopod of 3 segments, middle segment with a spine on outer margin, or leg 1 exopod of only 1 or 2 segments; body stout,
usually not vermiform ......................................................................................................................................................................48
47a(46a).
Legs 1 and 2, endopods of 2 and 1 segment, respectively ................................................................Parastenocarididae Parastenocaris
47b.
Legs 1 and 2, endopods of 3 and 2 segments, respectively .......................................................................Ameiridae Psammonitocrella
48a(46b)
Second antenna with basis (Fig. 3) ...................................................................................................................................................49
48b.
Second antenna with allobasis (Fig. 3)..............................................................................................................................................51
49a(48a).
Leg 5 unsegmented .....................................................................................................................................Tachidiidae Tachidius
49b.
Leg 5 of 2 distinct segments, i.e. exopod segment separated from baseoendopod ...................................................Ameiridae 50
50a(49b).
Caudal rami short, 1-2 times as broad as long; legs 2 – 4, 1 or more endopod 3-segmented..............................................................65
50b.
Caudal rami about 5 times longer than wide; legs 2 – 4, endopod with 2, 2, and 1 segments, respectively.....................Stygonitocrella
51a(48b).
Leg 1 with endopod of 2 or 3 segments ............................................................................................................................................52
51b.
Leg 1 with endopod of 1 short segment ..............................................................................................Huntemanniidae Huntemannia
52a(51a).
First antenna of 6 – 9 segments; leg 1 , endopod segment 1 with inner seta............................................................................53
52b.
First antenna of 5 segments; leg 1 , endopod segment 1 without inner seta ...................................Huntemanniidae Nannopus
53a(52a).
First antenna of 6 – 9 (usually 8) segments; leg 5 of 2 distinct segments, i.e., exopod separated from baseoendopod
...........................................................................................................................................................................Canthocamptidae 54
53b.
First antenna of 6 segments; leg 5 unsegmented ...............................................Canthocamptidae incertae sedis Cletocamptus
942
C. E. Williamson and J. W. Reid
54a(53).
Leg 1 , exopod of 3 segments ....................................................................................................................................................55
54b.
Leg 1 , exopod of 2 segments.................................................................................................................................Maraenobiotus
55a(54a).
Leg 1 , exopod segment 2 without inner seta.............................................................................................................................56
55b.
Leg 1 , exopod segment 2 with inner seta ..................................................................................................................................60
56a(55a).
Leg 1 , endopod of 2 segments ..................................................................................................................................................57
56b.
Leg 1 , endopod of 3 segments ....................................................................................................................Bryocamptus (in part)
57a(56a).
Leg 1 , endopod reaches from about midlength to slightly past end of exopod segment 3 .........................................................58
57b.
Leg 1 , endopod does not reach end of exopod segment 2 .......................................................................................Epactophanes
57c.
Leg 1 , endopod longer than exopod by half of endopod segment 3 or more ............................................................Paracamptus
58a(57a).
Legs 2 – 4 , exopod 2 without inner seta ....................................................................................................................................59
58b.
Legs 2 – 4 , exopod 2 with inner seta ...........................................................................................................Bryocamptus (in part)
59a(58a).
Leg 3 , exopod segment 3 with 1 inner seta; legs 2 – 4 , exopod segment 3 with all normal-sized setae and spines (i.e., inner
terminal seta not reduced) .......................................................................................................................................................Moraria
59b.
Leg 3 , exopod segment 3 with 2 inner setae; legs 2 – 4 , exopod segment 3, inner terminal seta tiny...................Gulcamptus
60a(55b).
Leg 1 , endopod of 3 segments and rostrum small to moderate in size, i.e., not extending past segment 1 of first antenna; first
antenna of 7 – 9 (usually 8) segments ............................................................................................................................................61
60b.
Leg 1 , endopod of 2 segments and rostrum small, not extending past segment 1 of first antenna; first antenna of 7 or 8 segments ................................................................................................................................................................Bryocamptus (in part)
60c.
Leg 1 , endopod of 2 or 3 segments and rostrum large, extending past end of segment 1 of first antenna; first antenna of 6 or
7 segments ............................................................................................................................................................................Mesochra
61a(60a).
Leg 5 , next outermost seta of baseoendopod not reduced, usually equal or longer than outermost seta; leg 4 , outer corner of
last segment of endopod with distinct spine or seta ..........................................................................................................................62
61b.
Leg 5 , next outermost seta of baseoendopod tiny; leg 4 , outer corner of last segment of endopod with spinous process
....................................................................................................................................................................................Canthocamptus
62a(61a).
Leg 1 , endopod segment 1 not reaching beyond end of exopod segment 2 ...............................................................................63
62b.
Leg 1 , endopod segment 1 reaching midlength of exopod segment 3, or longer ..............................................Attheyella (in part)
63a(62a).
Legs 2 – 4 , exopod segment 3 with 3 outer spines (total number of spines and setae 6, 7, and 6 or 7, respectively); leg 5 ,
baseoendopod with 5 or 6 (rarely 4) setae; leg 5 , baseoendopod usually with 2 setae ..................................................................64
63b.
Legs 2 – 4 , exopod segment 3 with 2 outer spines (total number of spines and setae 5, 6, and 6, respectively); leg 5 , baseoendopod with 3 – 4 setae; leg 5 , baseoendopod with no setae ...........................................................................................Elaphoidella
64a(63).
Legs 2 and 3 , endopods of 3 segments; or of 2 segments and leg 5 with 5 setae on baseoendopod; leg 2 , endopod segment 2
modified, bearing 1 or 2 knobs, spinous processes, or notches ...........................................................................Bryocamptus (in part)
64b
Legs 2 and 3 , endopods of 2 segments; leg 5 with 6 setae on baseoendopod; leg 2 , endopod unmodified (may have spines on
outer margin)..........................................................................................................................................................Attheyella (in part)
65a(50a).
Leg 2-4 , all endopods of 3 segments ................................................................................................................................Nitokra
65b.
Leg 2-4 , endopods of 3, 3 and 2 segments respectively ...........................................................................................Nitocrellopsis
LITERATURE CITED
Adrian, R. 1991. The feeding behavior of Cyclops kolensis and C.
vicinus (Crustacea, Copepoda). Verhandlungen der Internationale Vereinigung für Theoretische und Angewandte Limnologie 24:2852 – 2863.
Adrian, R. 1997. Calanoid – cyclopoid interactions: evidence from an
11-year field study in a eutrophic lake. Freshwater Biology
38:315 – 326.
Adrian, R., Frost, T. M. 1992. Comparative feeding ecology of
Tropocyclops prasinus mexicanus (Copepoda, Cyclopoida).
Journal of Plankton Research 14:1369 – 1382.
Allan, J. D. 1976. Life history patterns in zooplankton. American
Naturalist 110:165 – 180.
Andersen, T., Hessen, D. O. 1991. Carbon, nitrogen, and phosphorus content of freshwater zooplankton. Limnology and
Oceanography 36:807 – 814.
Anderson, R. S. 1970. Predator – prey relationships and predation
rates for crustacean zooplankters from some lakes in western
Canada. Canadian Journal of Zoology 48:1229 – 1240.
Arts, M. T., Evans, M. S. 1991. Optical – digital measurements of energy reserves in calanoid copepods: Intersegmental distribution
and seasonal patterns. Limnology and Oceanography
36:289 – 298.
22. Copepoda
Barszcz, C. A., Yevich, P. P. 1976. Preparation of copepods for
histopathological examinations. Transactions of the American
Microscopical Society 95:104 – 108.
Batchelder, H. P. 1986. A staining technique for determining copepod
gonad maturation: application to Metridia pacifica from the northern Pacific Ocean. Journal of Crustacean Biology 6: 227– 231.
Blades, P. I., Youngbluth, M. J. 1980. Morphological, physiological,
and behavioral aspects of mating in calanoid copepods. In: Kerfoot, W. C., Ed., Evolution and ecology of zooplankton communities. Univ. Press of New England, Hanover, NH, pp. 39 – 51.
Blais, J. M., Maly, E. J. 1993. Differential predation by Chaoborus
americanus on males and females of two species of Diaptomus.
Canadian Journal of Fisheries and Aquatic Sciences
50:410 – 415.
Bogdan, K.G., Gilbert, J.J. 1984. Body size and food size in freshwater zooplankton. Proceedings of the National Academy of Sciences 81:6427 – 6431.
Bourguet, J.-P. 1986. Contribution àl’étude de Cletocamptus retrogressus Schmankewitch, 1875 (Copepoda, Harpacticoida) II.
Développement larvaire — stades naupliens. Crustaceana
51:113 – 122
Bowers, J. A. 1986. Phosphorus regeneration by the predatory copepod Diacyclops thomasi. Canadian Journal of Fisheries and
Aquatic Sciences 43:361 – 365.
Bowman, T. E., Abele, L. G. 1982. Classification of the recent Crustacea. Pages 1 – 25, in: Bliss, D. E., Ed. The biology of Crustacea. Vol. 1, Abele, L. G., Ed., Systematics, the fossil record,
and biogeography, Academic Press, New York.
Brandl, Z. 1998. Feeding strategies of planktonic cyclopoids in lacustrine ecosystems. Journal of Marine Systems 15:87 – 95.
Brandl, Z., Fernando, C. H. 1979. The impact of predation by the
copepod Mesocyclops edax (Forbes) on zooplankton in three
lakes in Ontario, Canada. Canadian Journal of Zoology
57:940 – 942.
Brandl, Z., Fernando, C. H. 1986. Feeding and food consumption by
Mesocyclops edax. Syllogeus 58:254 – 258.
Brown, M. D., Kay, B. H., Greenwood, J. G. 1991. The predation efficiency of north-eastern Australian Mesocyclops (Copepoda:
Cyclopoida) on mosquito larvae. Bulletin of the Plankton Society of Japan, Special Vol.:329 – 338.
Bulkowski, L., Krise, W. F., Kraus, K. A. 1985. Purification of
Cyclops cultures by pH shock (Copepoda). Crustaceana
48:179 – 182.
Buskey, E. J. 1998. Components of mating behavior in planktonic
copepods. Journal of Marine Systems. 15:13 – 21.
Bunting, D. L. 1973. The Cladocera and Copepoda of Tennessee. II.
Cyclopoid copepods. Journal of the Tennessee Academy of Science 48:138 – 141.
Burgi, H.-R., Elser, J. J., Richards, R. C., Goldman, C. R. 1993. Zooplankton patchiness in Lake Tahoe and Castle Lake U.S.A. Verhandlungen der Internationale Vereinigung für Theoretische
und Angewandte Limnologie 25:378 – 382.
Burns, C. W., Gilbert, J. J. 1993. Predation on ciliates by freshwater
calanoid copepods: rates of predation and relative vulnerabilities of prey. Freshwater Biology 30:377 – 393.
Burns, C. W., Schallenberg, M. 1996. Relative impacts of copepods,
cladocerans and nutrients on the microbial food web of a
mesotrophic lake. Journal of Plankton Research 18:683 – 714.
Byron, E. R., Folt, C. L., Goldman, C. R. 1984. Copepod and cladoceran success in an oligotrophic lake. Journal of Plankton Research 6:45 – 65.
Byron, E. R., Whitman, P. T., Goldman, C. R. 1983. Observations of
copepod swarms in Lake Tahoe. Limnology and Oceanography
28:378 – 382.
Carter, J. C. H., Goudie, K. A. 1986. Diel vertical migrations and
943
horizontal distributions of Limnocalanus macrurus and Senecella calanoides (Copepoda, Calanoida) in lakes of southern
Ontario in relation to planktivorous fish. Canadian Journal of
Fisheries and Aquatic Sciences 43:2508 – 2514.
Carter, J. C. H., Sprules, W. G., Dadswell, M. J., Roff, J. C. 1983.
Factors governing geographical variation in body size of Diaptomus minutus (Copepoda, Calanoida). Canadian Journal of
Fisheries and Aquatic Sciences 40:1303 – 1307.
Chaston, I. 1969. Anaerobiosis in Cyclops varicans. Limnology and
Oceanography 14:298 – 301.
Chen, C. Y., Folt, C. L., Cook, S. 1997. The potential for hybridization in freshwater copepods. Oecologia 111:557 – 564.
Chengalath, R., Shih, C-t. 1994. Littoral freshwater copepods of
northwestern North America: northern British Columbia. Verhandlungen der Internationale Vereinigung für Theoretische
und Angewandte Limnologie 25:2421 – 2431.
Chow-Fraser, P., Maly, E. J. 1988. Aspects of mating, reproduction,
and co-occurrence in three freshwater calanoid copepods.
Freshwater Biology 19:95 – 108.
Chow-Fraser, P., Maly, E. J. 1991. Factors governing clutch size in
two species of Diaptomus (Copepoda:Calanoida). Canadian
Journal of Fisheries and Aquatic Sciences 48:364 – 370.
Chow-Fraser, P., Wong, C. K. 1986. Dietary change during development in the freshwater calanoid copepod Epischura lacustris
Forbes. Canadian Journal of Fisheries and Aquatic Sciences
43:938 – 944.
Clamp, J. C. (Compiler), Adams, W. F., Reid, J. W., Taylor, A. Y.,
Cooper, J. E., McGrath, C., Williams, D. J., DeMont, D. J.,
McLarney, W. O., Mottesi, G., Alderman, J. 1999. A report on
the conservation status of North Carolina’s freshwater and terrestrial crustacean fauna. Scientific Council on Freshwater and
Terrestrial Crustaceans, North Carolina Wildlife Resources
Commission, Raleigh. 92 p.
Cole, G. A. 1959. A summary of our knowledge of Kentucky crustaceans. Transactions of the Kentucky Academy of Science
20:66 – 81.
Cole, G. A. 1961. Some calanoid copepods from Arizona with notes
on congeneric occurrences of Diaptomus species. Limnology
and Oceanography 6:432 – 442.
Cole, G. A. 1963. Calanoid copepods from some old Arizona collections. Journal of the Arizona Academy of Science 2:176 – 183.
Cole, G.A. 1966. Contrasts among calanoid copepods from permanent and temporary ponds in Arizona. American Midland Naturalist 76:351 – 368.
Comita, G. W., McNett, S. J. 1976. The postembryonic developmental instars of Diaptomus oregonensis Lilljeborg, 1889 (Copepoda). Crustaceana 30:123 – 163.
Comita, G. W., Tommerdahl, D. M. 1960. The postembryonic developmental instars of Diaptomus siciloides Lilljeborg. Journal of
Morphology 107:297 – 355.
Confer, J. L., Blades, P. I. 1975. Omnivorous zooplankton and planktivorous fish. Limnology and Oceanography 20:571 – 579.
Confer, J. L., Cooley, J. M. 1977. Copepod instar survival and predation by zooplankton. Journal of the Fisheries Research Board of
Canada 34:703 – 706.
Confer, J. L., Kaaret, T., Likens, G. E. 1983. Zooplankton diversity
and biomass in recently acidified lakes. Canadian Journal of
Fisheries and Aquatic Sciences 40:36 – 42.
Coull, B. C. 1977. Marine flora and fauna of the northeastern United
States. Copepoda: Harpacticoida. NOAA Technical Report
NMFS Circular 399, 48 pp.
Czaika, S. C. 1982. Identification of nauplii N1 – N6 and copepodids
CI – CVI of the Great Lakes calanoid and cyclopoid copepods
(Calanoida, Cyclopoida, Copepoda). Journal of Great Lakes
Research 8:439 – 469.
944
C. E. Williamson and J. W. Reid
Czaika, S. C., Robertson, A. 1968. Identification of the copepodids
of the Great Lakes species of Diaptomus (Calanoida, Copepoda). Proceedings of the 11th Conference on Great Lakes Research 17:39 – 60.
DeMott, W. R. 1986. The role of taste in food selection by freshwater zooplankton. Oecologia 69:334 – 340.
DeMott, W. R. 1988. Discrimination between algae and artificial
particles by freshwater and marine copepods. Limnology and
Oceanography 33:397 – 408.
DeMott, W. R. 1989. Discrimination between algae and detritus by
freshwater and marine zooplankton. Bulletin of Marine Science
43:486 – 499.
DeMott, W. R. 1995a. Food selection by calanoid copepods in response to between-lake variation in food abundance. Freshwater Biology 33:171 – 180.
DeMott, W. R. 1995b. Optimal foraging by a suspension-feeding
copepod: responses to short-term and seasonal variation in food
resources. Oecologia 103:230 – 240.
DeMott, W. R., Moxter, F. 1991. Foraging on cyanobacteria by copepods: responses to chemical defenses and resource abundance.
Ecology 72:1820 – 1834.
DeMott, W. R., Zhang, Q.-X., Carmichael, W. W. 1991. Effects of
toxic cyanobacteria and purified toxins on the survival and
feeding of a copepod and three species of Daphnia. Limnology
and Oceanography 36:1346 – 1357.
De Stasio, B. T. Jr. 1990. The role of dormancy and emergence patterns in the dynamics of a freshwater zooplankton community.
Limnology and Oceanography 35:1079 – 1090.
De Stasio, B. T. Jr. 1993. Diel vertical and horizontal migration by
zooplankton:population budgets and the diurnal deficit. Bulletin
of Marine Science 53:44 – 64.
Dobrzykowski, A. E., Wyngaard, G. A. 1993. Phenology of dormancy in a Virginia population of Mesocyclops edax (Crustacea: Copepoda). Hydrobiologia 250:167 – 171.
Dodson, S. I., Egger, D. L. 1980. Selective feeding of red phalaropes
on zooplankton of arctic ponds. Ecology 61:755 – 763.
Donald, D. B., Anderson, R. S., Mayhood, D. W. 1994. Coexistence
of fish and large Hesperodiaptomus species (Crustacea:
Calanoida) in subalpine and alpine lakes. Canadian Journal of
Zoology 72:259 – 261.
Dorward, H. M., Wyngaard, G. A. 1997. Variability and pattern of
chromatin diminution in the freshwater Cyclopidae (Crustacea:
Copepoda). Archiv für Hydrobiologie, Supplement 107:
447 – 465.
Dressel, D. M., Heinle, D. R., Grote, M. C. 1972. Vital staining to
sort dead and live copepods. Chesapeake Science 13:156 – 159.
Dürbaum, J. 1995. Discovery of postcopulatory mate guarding in
Copepoda Harpacticoida (Crustacea). Marine Biology
123:81 – 88.
Dürbaum, J. 1997. Precopulatory mate guarding and mating in
Tachidius discipes (Copepoda:Harpacticoida). Contributions to
Zoology 66:201 – 214.
Dussart, B. H., Defaye, D. 1995. Introduction to the Copepoda.
Guides to the Identification of the Microinvertebrates of the
Continental Waters of the World. Dumont, H. J. (Ed.) 7:1 – 277.
Edmondson, W. T. 1960. Reproductive rates of rotifers in natural populations. Memorie dell’Istituto Italiano di Idrobiologia 12:21 – 77.
Edmondson, W. T. 1964. The rate of egg production by rotifers and
copepods in natural populations as controlled by food and temperature. Verhandlungen der Internationale Vereinigung für
Theoretische und Angewandte Limnologie 15:673 – 675.
Edmondson, W. T. 1985. Reciprocal changes in abundance of Diaptomus and Daphnia in Lake Washington. Archiv für Hydrobiologie, Beihefte Ergebnisse der Limnologie 21:475 – 481.
Edmondson, W. T., Comita, G. W., Anderson, G. C. 1962. Reproductive rate of copepods in nature and its relation to phytoplankton population. Ecology 43:625 – 634.
Einsle, U. 1996. Copepoda:Cyclopoida. Genera Cyclops, Megacyclops, Acanthocyclops. Guides to the Identification of the
Microinvertebrates of the Continental Waters of the World. Dumont, H.J. (Ed.) 10:1 – 83.
Einsle, U. 1996. Cyclops heberti n. sp. and Cyclops singularis n. sp.,
two new species within the genus Cyclops (‘strenuus-subgroup’)
(Crustacea: Copepoda) from ephemeral ponds in southern
Germany. Hydrobiologia 319:167 – 177.
Elgmork, K. 1967. Ecological aspects of diapause in copepods. Symposium Series of the Marine Biological Association of India
2(3): 947 – 954 and 1 plate.
Elgmork, K., Nilssen, J. P. 1978. Equivalence of copepod and insect
diapause. Verhandlungen der Internationale Vereinigung für
Theoretische und Angewandte Limnologie 20:2511 – 2517.
Elmore, J. L. 1982. The influence of food concentration and container volume on life history parameters of Diaptomus dorsalis
Marsh from subtropical Florida. Hydrobiologia 89:215 – 223.
Elmore, J. L. 1983. Factors influencing Diaptomus distributions: An
experimental study in subtropical Florida. Limnology and
Oceanography 28:522 – 532.
Elofsson, R. 1971. The ultrastructure of a chemoreceptor organ in
the head of copepod crustaceans. Acta Zoologica 52:299 – 315.
Fahrenbach, W. H. 1962. The biology of a harpacticoid copepod.
Cellule 62:303 – 376.
Feifarek, B. P., Wyngaard, G. A., Allan, J. D. 1983. The cost of reproduction in a freshwater copepod. Oecologia 56:166 – 168.
Feller, R. J. 1980. Development of the sand-dwelling meiobenthic
harpacticoid copepod Huntemannia jadensis Poppe in the laboratory. Journal of Experimental Marine Biology and Ecology
46:1 – 15.
Fiers, F., Reid, J. W., Iliffe, T. M., Suárez-Morales, E. 1996. New hypogean cyclopoid copepods (Crustacea) from the Yucatan
Peninsula, Mexico. Contributions to Zoology 66:65 – 102.
Fischer, J. M., Frost, T. M. 1997. Indirect effects of lake acidification
on Chaoborus population dynamics: the role of food limitation
and predation. Canadian Journal of Fisheries and Aquatic Sciences 54:637 – 646.
Fischer, J. M., Moore, M. V. 1993. Juvenile survival of a planktonic
insect:effects of food limitation and predation. Freshwater Biology 30:35 – 45.
Fleminger, A. 1973. Pattern, number, variability, and taxonomic significance of integumental organs (sensilla and glandular pores)
in the genus Eucalanus (Copepoda, Calanoida). Fishery Bulletin
71:965 – 1010.
Flemming, J. M., Coughlan, J. 1978. Preservation of vitally stained
zooplankton for live / dead sorting. Estuaries 1:135 – 137.
Folt, C. L., Goldman, C. R. 1981. Allelopathy between zooplankton: A
mechanism for interference competition. Science 213:1133 – 1135.
Frenzel, P. 1980. Die Populationsdynamik von Canthocamptus
staphylinus (Jurine) (Copepoda, Harpacticoida) im Litoral des
Bodensees. Crustaceana 39:282 – 286.
Friedman, M. M. 1980. Comparative morphology and functional significance of copepod receptors and oral structures. in: Kerfoot,
W. C. (Ed.), Evolution and ecology of zooplankton communities.
Uni. Press of New England, Hanover, NH, Pages 185 – 197.
Friedman, M. M., Strickler, J. R. 1975. Chemoreceptors and feeding
in calanoid copepods (Arthropoda:Crustacea). Proceedings of
the National Academy of Sciences of the U.S.A. 72:4185 – 4188.
Fryer, G. 1957a. The feeding mechanism of some freshwater cyclopoid copepods. Proceedings of the Zoological Society of
London 129:1 – 25.
22. Copepoda
Fryer, G. 1957b. The food of some freshwater cyclopoid copepods
and its ecological significance. Journal of Animal Ecology
26:263 – 286.
Gabriel, W. 1985. Overcoming food limitation by cannibalism: a
model study on cyclopoids. Archiv für Hydrobiologie, Beihefte
Ergebnisse der Limnologie 21:373 – 381.
Gannon, J. E. 1971. Two counting cells for the enumeration of zooplankton micro-Crustacea. Transactions of the American Microscopical Society 90:486 – 490.
Gannon, J. E., Gannon, S. A. 1975. Observations on the narcotization of crustacean zooplankton. Crustaceana 28:220 – 224.
Gauld, D. T. 1951. The grazing rate of planktonic copepods. Journal
of the Marine Biological Association of the United Kingdom
29:695 – 706.
Gee, J. M., Warwick, R. M. 1984. Preliminary observations on the
metabolic and reproductive strategies of harpacticoid copepods
from an intertidal sandflat. Hydrobiologia 118:29 – 37.
Gehrs, C. W. 1974. Horizontal distribution and abundance of Diaptomus clavipes Schacht in relation to Potamogeton foliosus in a
pond and under experimental conditions. Limnology and
Oceanography 19:100 – 104.
Gerritsen, J. 1978. Instar-specific swimming patterns and predation
of planktonic copepods. Verhandlungen der Internationale Vereinigung für Theoretische und Angewandte Limnologie
20:2531 – 2536.
Gilbert, J. J., Williamson, C. E. 1978. Predator – prey behavior and
its effect on rotifer survival in associations of Mesocyclops
edax, Asplanchna girodi, Polyarthra vulgaris, and Keratella
cochlearis. Oecologia 37:13 – 22.
Gilbert, J. J., Williamson, C. E. 1983. Sexual dimorphism in zooplankton (Copepoda, Cladocera, and Rotifera). Annual Review
of Ecology and Systematics 14:1 – 33.
Glatzel, T., Schminke, H. K. 1996. Mating behaviour of the groundwater copepod Parastenocaris phyllura Kiefer, 1938 (Copepoda:Harpacticoida). Contributions to Zoology 66:103 – 108.
Gliwicz, Z. M. 1994. Retarded growth of cladoceran zooplankton in
the presence of a copepod predator. Oecologia 97:458 – 461.
Gliwicz, Z. M., Lampert, W. 1994. Clutch-size variability in
Daphnia: Body-size related effects of egg predation by cyclopoid copepods. Limnology and Oceanography 39:479 – 485.
Gliwicz, Z. M., Stibor, H. 1993. Egg predation by copepods in
Daphnia brood cavities. Oecologia 95:295 – 298.
Gliwicz, Z. M., Umana, G. 1994. Cladoceran body size and vulnerability to copepod predation. Limnology and Oceanography
39:419 – 424.
Gophen, M. 1978. Errors in the estimation of recruitment of early
stages of Mesocyclops leuckarti (Claus) caused by the diurnal
periodicity of egg-production. Hydrobiologia 57:59 – 64.
Gophen, M. 1979. Mating process in Mesocyclops leuckarti (Crustacea:Copepoda). Israel Journal of Zoology 28:163 – 166.
Gophen, M., Harris, R. P. 1981. Visual predation by a marine cyclopoid copepod, Corycaeus anglicus. Journal of the Marine Biological Association of the United Kingdom 61:391 – 399.
Gould, D. T. 1957. A peritrophic membrane in calanoid copepods.
Nature 179:325 – 326.
Grad, G., Maly, E. J. 1988. Sex size ratios and their influence on
mating success in a calanoid copepod. Limnology and Oceanography 33:1629 – 1634.
Grad, G., Maly, E. J. 1992. Further observations relating sex size ratios to mating success in calanoid copepods. Journal of Plankon
Research 14: 903 – 913
Guma’a, S. A. 1978. The food and feeding habits of young perch,
Perca fluviatilis, in Windermere. Freshwater Biology 8:
177 – 187.
945
Hairston, N. G. Jr. 1976. Photoprotection by carotenoid pigments in
the copepod Diaptomus nevadensis. Proceedings of the National Academy of Sciences of the U.S.A. 73:971 – 974.
Hairston, N. G. Jr. 1979a. The adaptive significance of color polymorphism in two species of Diaptomus (Copepoda). Limnology
and Oceanography 24:15 – 37.
Hairston, N. G. Jr. 1979b. The relationship between pigmentation
and reproduction in two species of Diaptomus (Copepoda).
Limnology and Oceanography 24:38 – 44.
Hairston, N. G. Jr. 1979c. The effect of temperature on carotenoid
photoprotection in the copepod Diaptomus nevadensis. Comparative Biochemistry and Physiology 62:445 – 448.
Hairston, N. G. Jr. 1996. Zooplankton egg banks as biotic reservoirs
in changing environments. Limnology and Oceanography
41:1087 – 1092.
Hairston, N. G. Jr., Olds, E. J. 1984. Population differences in the
timing of diapause: adaptation in a spatially heterogeneous environment. Oecologiaphy 61:42 – 48.
Hairston, N. G., Jr., Twombly, S. 1985. Obtaining life table data
from cohort analyses: A critique of current methods. Limnology
and Oceanography. 30:886 – 893.
Hairston, N. G. Jr., Van Brunt, R. A. 1994. Diapause dynamics of
two diaptomid copepod species in a large lake. Hydrobiologia
292 / 293:209 – 218.
Hairston, N. G. Jr., Van Brunt, R. A., Kearns, C. M., Engstrom, D.
R. 1995. Age and survivorship of diapausing eggs in a sediment
egg bank. Ecology 76:1706 – 1711.
Hairston, N. G. Jr., Walton, W. E. 1986. Rapid evolution of a life
history trait. Proceedings of the National Academy of Sciences
of the U.S.A. 83:4831 – 4833.
Hairston, N. G. Jr., Walton, W. E., Li, K. T. 1983. The causes and
consequences of sex-specific mortality in a freshwater copepod.
Limnology and Oceanography 28:935 – 947.
Hansen, A.-M., Santer, B. 1995. The influence of food resources on
the development, survival and reproduction of the two cyclopoid copepods: Cyclops vicinus and Mesocyclops leuckarti.
Journal of Plankton Research 17:631 – 646.
Hardy, A. 1970. The Open Sea. The World of Plankton. Collins,
London, 335 p.
Harris, M. J. 1978. Copepoda of northern Mississippi with a description of a new subspecies. Tulane Studies in Zoology and
Botany 20:27 – 34.
Hart, R. C., Santer, B. 1994. Nutritional suitability of some uni-algal
diets for freshwater calanoids: unexpected inadequacies of commonly used edible greens and others. Freshwater Biology
31:109 – 116.
Hartig, J. H., Jude, D. J., Evans, M. S. 1982. Cyclopoid predation on
Lake Michigan fish larvae. Canadian Journal of Fisheries and
Aquatic Sciences 39:1563 – 1568.
Hartmann, H. J., Taleb, H., Aleya, L., Lair, N. 1993. Predation on
ciliates by the suspension-feeding calanoid copepod Acanthodiaptomus denticornis. Canadian Journal of Fisheries and Aquatic
Sciences 50:1382 – 1393.
Hebert, P. D. N. 1985. Ecology of the dominant copepod species at a
Low Arctic site. Canadian Journal of Zoology 63:1138 – 1147.
Hebert, P. D. N., Good, A. G., Mort, M. A. 1980. Induced swarming
in the predatory copepod Heterocope septentrionalis. Limnology and Oceanography 25:747 – 750.
Herbst, H.-V. 1986. Beschreibung des Thermocyclops hastatus antillensis n. ssp. mit einem Bestimmungschlüssel für die Gattung Thermocyclops Kiefer, 1927. Bijdragen tot de Dierkunde 56:165 – 180.
Hicks, G. R. F., Coull, B. C. 1983. The ecology of marine meiobenthic harpacticoid copepods. Annual Review of Oceanography
and Marine Biology 21:67 – 175.
946
C. E. Williamson and J. W. Reid
Hopp, U., Maier, G., Bleher, R. 1997. Reproduction and adult
longevity of five species of planktonic cyclopoid copepods
reared on different diets: a comparative study. Freshwater Biology 38:289 – 300.
Hosfeld, B. 1996. The relationship between the rostrum and the organ
of Bellonci in copepods: an ultrastructural study of the rostrum
of Canuella perplexa. Zoologischer Anzeiger 234:175 – 190.
Hosfeld, B., Schminke, H. K. 1997. The ultrastructure of ionocytes
from osmoregulatory integumental windows of Parastenocaris
vicesima (Crustacea, Copepoda, Harpacticoida). Archiv für Hydrobiologie 139:389 – 400.
Hudson, P. L., Reid, J. W., Lesko, L. T., Selgeby, J. H. 1998. Cyclopoid and harpacticoid copepods of the Laurentian Great
Lakes. Bulletin of the Ohio Biological Survey, New Series. 12:
vii, 1 – 50 pp.
Huq, A., Small, E. B., West, P. A., Huq, M. I., Rahman, R., Colwell,
R. R. 1983. Ecological relationships between Vibrio cholerae
and planktonic crustacean copepods. Applied and Environmental Microbiology 45:275 – 283.
Hutchinson, G. E. 1967. A Treatise on Limnology, Vol. 2. Wiley,
New York.
Huys, R., Boxshall, G. A. 1991. Copepod Evolution. The Ray Society, London.
Huys, R., Gee, J. M., Moore, C. G., Hamond, R. 1996. Marine and
brackish water harpacticoid copepods. Part 1. Keys and notes
for identification of the species. Synopses of the British Fauna,
New Series 51:1 – 352.
Ianora, A., Miralto, A., Buttino, I., Romano, G. 1999. First evidence
of some dinoflagellates reducing male copepod fertilization capacity. Limnology and Oceanography 44:147 – 153.
Jamieson, C. D. 1980. Observations on the effect of diet and temperature on rate of development of Mesocyclops leuckarti (Claus)
(Copepoda, Cyclopoida). Crustaceana 38:145 – 154.
Janetzky, W., Martínez Arbizu, P., Reid, J. W. 1996. Attheyella (Canthosella) mervini sp. n. (Canthocamptidae, Harpacticoida) from
Jamaican bromeliads. Hydrobiologia 339:123 – 135.
Jersabek, C. D., Schabetsberger, R. 1995. Resting egg production and
oviducal cycling in two sympatric species of alpine diaptomids
(Copepoda: Calanoida) in relation to temperature and food
availability. Journal of Plankton Research 17:2049 – 2078.
Karaytug, S. 1999. Cyclopoida — Genera Paracyclops, Ochridacyclops and key to the Eucyclopina. Guides to the Identification of
the Microinvertebrates of the Continental Waters of the World.
Dumont, H. J. (Ed.) 14: 1 – 217
Katona, S. K. 1971. The developmental stages of Eurytemora affinis
(Poppe, 1880) (Copepoda, Calanoida) raised in laboratory cultures, including a comparison with the larvae of Eurytemora
americana Williams, 1906, and Eurytemora herdmani Thompson and Scott, 1897. Crustaceana 21:5 – 20.
Kerfoot, W. C. 1978. Combat between predatory copepods and their
prey: Cyclops, Epischura, and Bosmina. Limnology and
Oceanography 23:1089 – 1102.
Kerfoot, W. C., DeMott, W. R. 1984. Food web dynamics: Dependent chains and vaulting. Pages 347 – 382 in: Meyers, D. G.,
Strickler, J. R. Ed., Trophic interactions within aquatic ecosystems. American Association for the Advancement of Science Selected Symposium 85. Westview Press Inc., Boulder, CO.
Koehl, M. A. R., Strickler, J. R. 1981. Copepod feeding currents:
Food capture at low Reynolds number. Limnology and
Oceanography 26:1062 – 1073.
Lampert, W., Muck, P. 1985. Multiple aspects of food limitation in
zooplankton communities: the Daphnia-Eudiaptomus example.
Archiv für Hydrobiologie, Beihefte Ergebnisse der Limnologie
21:311 – 322.
Landry, M. R., Hassett, R. P. 1982. Estimating the grazing impact of
marine micro-zooplankton. Marine Biology 67:283 – 288.
Lang, K. 1948. Monographie der Harpacticiden. Vols. I, II. H. Ohlsson, Lund.
Lang, K. 1965. Copepoda Harpacticoida from the Californian Pacific
coast. Kunglige Svenska Vetenskapsakademiens Handlingar,
N.S., 10(2):1 – 560 5 plates.
Leech, D. M., Wyngaard, G. A. 1996. Timing of chromatin diminution in the free-living, freshwater Cyclopidae (Crustacea:Copepoda). Journal of Crustacean Biology 16:496 – 500.
Legier-Visser, M. F., Mitchell, J. G., Okubo, A., Fuhrman, J. A. 1986.
Mechanoreception in calanoid copepods. A mechanism for prey
detection. Marine Biology 90:529 – 535.
Lehman, J. T. 1980. Release and cycling of nutrients between planktonic algae and herbivores. Limnology and Oceanography
25:620 – 632.
Lehman, P. S., Reid, J. W. 1993. Phyllognathopus viguieri (Crustacea: Harpacticoida), a predaceous copepod of phytoparasitic,
entomopathogenic, and free-living nematodes. Soil and Crop
Science Society of Florida Proceedings 52:78 – 82.
Lescher-Moutoué, F. 1996. Seasonal variations in size and morphology of Acanthocyclops robustus (Copepoda Cyclopoida). Journal of Plankton Research 18:907 – 922.
Lewis, W. M. Jr., Saunders, J. F. III. 1979. Two new integrating samplers for zooplankton, phytoplankton, and water chemistry.
Archiv für Hydrobiologie 85:244 – 249.
Liebig, J. R., Vanderploeg, H. A. 1995. Vulnerability of Dreissena
polymorpha larvae to predation by Great Lakes calanoid copepods: The importance of the bivalve shell. Journal of Great
Lakes Research 21:353 – 358.
Lohner, L. M., Hairston, Jr., N. G., Schaffner, W. R. 1990. A method
for distinguishing subitaneous and diapausing eggs in preserved
samples of the calanoid copepod genus Diaptomus. Limnology
and Oceanography 35:763 – 767.
Lonsdale, D. J., Frey, M. A., Snell, T. W. 1998. The role of chemical
signals in copepod reproduction. Journal of Marine Systems
15:1 – 12.
Luecke, C., Litt, A. H. 1987. Effects of predation by Chaoborus flavicans on crustacean zooplankton of Lake Lenore, Washington.
Freshwater Biology 18:185 – 192.
Luecke, C., O’Brien, W. J. 1981. Phototoxicity and fish predation:
Selective factors in color morphs in Heterocope. Limnology and
Oceanography 26:454 – 460.
Luecke, C., O’Brien, W. J. 1983. The effect of Heterocope predation
on zooplankton communities in arctic ponds. Limnology and
Oceanography 28:367 – 377.
Maier, G. 1995. Mating frequency and interspecific matings in some
freshwater cyclopoid copepods. Oecologia 101:245 – 250.
Main, T. M., Dobberfuhl, D. R., Elser, J. J. 1997. N:P stoichiometry
and ontogeny of crustacean zooplankton: a test of the growth
rate hypothesis. Limnology and Oceanography 42:1474 – 1478.
Makarewicz, J. C., Likens, G. E. 1979. Structure and function of the
zooplankton community of Mirror Lake, New Hampshire. Ecological Monographs 49:109 – 127.
Maly, E. J. 1970. The influence of predation on the adult sex ratios
of two copepod species. Limnology and Oceanography
15:566 – 573.
Maly, E. J. 1973. Density, size, and clutch of two high altitude diaptomid copepods. Limnology and Oceanography 18:840 – 848.
Maly, E. J. 1976. Resource overlap between co-occurring copepods:
effects of predation and environmental fluctuation. Canadian
Journal of Zoology 54:933 – 940.
Maly, E. J., Maly, M. P. 1974. Dietary differences between two cooccurring calanoid copepod species. Oecologia 17:325 – 333.
22. Copepoda
Mantel, L. H., Farmer, L. L. 1983. Osmotic and ionic regulation. In:
Bliss, D. E. Ed., The Biology of Crustacea, Vol.5: Mantel, L. H.
Ed., Internal anatomy and physiological regulation. Academic
Press, New York, Pages 53 – 162.
Marshall, S. M., Orr, A. P. 1952. On the biology of Calanus finmarchicus. VII. Factors affecting egg production. Journal of the
Marine Biological Association of the United Kingdom
30:527 – 547.
Marten, G. G., Bordes, E. S., Nguyen, M. 1994. Use of cyclopoid
copepods for mosquito control. Hydrobiologia 292 / 293:
491 – 496.
Meyer, J. S., Ingersoll, C. G., McDonald, L. L., Boyce, M. S. 1986.
Estimating uncertainty in population growth rates: jackknife vs
bootstrap techniques. Ecology 67:1156 – 1166.
Monakov, A. V. 1976. Feeding and food interrelationships in freshwater copepods. Nauka Press, Leningrad, 170 p. [in Russian].
Moore, M. V., Folt, C. L., Stemberger, R. S. 1996. Consequences of
elevated temperatures for zooplankton assemblages in temperate lakes. Archiv für Hydrobiologie 135:289 – 319.
Morris, D. P., Zagarese, H., Williamson, C. E., Balseiro, E. G., Hargreaves, B. R., Modenutti, B., Moeller, R. E., Queimalinos, C.
1995. The attenuation of solar UV radiation in lakes and the
role of dissolved organic carbon. Limnology and Oceanography
40:1381 – 1391.
Musko, I. B. 1983. The structure of the alimentary canal of two
freshwater copepods of different feeding habits studied by light
microscope. Crustaceana 45:38 – 47.
Naess, T., Nilssen, J. P. 1991. Diapausing fertilized adults: A new
pattern of copepod life cycle. Oecologia 86:368 – 371.
Neill, W. E. 1990. Induced vertical migration in copepods as a defence against invertebrate predation. Nature 345:524 – 526.
Neill, W. E. 1992. Population variation in the ontogeny of predator
induced vertical migration of copepods. Nature 356:54 – 57.
Nie, H. W. de, Vijverberg, J. 1985. The accuracy of population density estimates of copepods and cladocerans, using data from
Tjeukemeer (The Netherlands) as an example. Hydrobiologia
124:3 – 11.
O’Brien, W. J., Schmidt, D. 1979. Arctic Bosmina morphology and
copepod predation. Limnology and Oceanography 24:564 – 568.
O’Doherty, E. C. 1985. Stream-dwelling copepods: Their life history
and ecological significance. Limnology and Oceanography
30:554 – 564.
O’Doherty, E. C. 1988. The ecology of meiofauna in an Appalachian
headwater stream. Ph.D. Dissertation, University of Georgia,
Athens, 113 p.
Orsi, J. J., Bowman, T. E., Marelli, D. C., Hutchinson, A. 1983.
Recent introduction of the planktonic calanoid copepod
Sinocalanus doerrii (Centropagidae) from mainland China to
the Sacramento-San Joaquin Estuary of California. Journal of
Plankton Research 5:357 – 375.
Pace, M. L. 1986. An empirical analysis of zooplankton community
size structure across lake trophic gradients. Limnology and
Oceanography 31:45 – 55.
Paloheimo, J. 1974. Calculation of instantaneous birth rate. Limnology and Oceanography 19:692 – 694.
Parker, B. R., Wilhelm, F. M., Schindler, D. W. 1996. Recovery of
Hesperodiaptomus arcticus populations from diapausing eggs
following elimination by stocked salmonids. Canadian Journal
of Zoology 74:1292 – 1297.
Paul, A. J., Leavitt, P. R., Schindler, D. W., Hardie, A. K. 1995. Direct and indirect effects of predation by a calanoid copepod
(subgenus:Hesperodiaptomus) and of nutrients in a fishless
alpine lake. Canadian Journal of Fisheries and Aquatic Sciences
52:2628 – 2638.
947
Paul, A. J., Schindler, D. W. 1994. Regulation of rotifers by predatory calanoid copepods (subgenus Hesperodiaptomus) in lakes
of the Canadian Rocky Mountains. Canadian Journal of Fisheries and Aquatic Sciences 51:2520 – 2528.
Peacock, A. H. 1982. Responses of Cyclops bicuspidatus thomasi to
alterations in food and predators. Canadian Journal of Zoology
60:1446 – 1462.
Peacock, A. H., Smyly, W. J. P. 1983. Experimental studies on the
factors limiting Tropocyclops prasinus (Fischer) 1860 in an oligotrophic lake. Canadian Journal of Zoology 61:250 – 265.
Pennak, R. W. 1989. Fresh-water Invertebrates of the United States.
Protozoa to Mollusca. 3rd ed. Wiley, New York.
Perlmutter, D. G., Meyer, J. L. 1991. The impact of a streamdwelling harpacticoid copepod upon detritally associated bacteria. Ecology 72:2170 – 2180.
Pinel-Alloul, B., Lamoureux, J. 1988a. Développement post-embryonnaire du copépode calanoïde Diaptomus (Aglaodiaptomus)
leptopus S. A. Forbes, 1882. I. Phase nauplienne. Crustaceana
54:69 – 84.
Pinel-Alloul, B., Lamoureux, J. 1988b. Développement post-embryonnaire du copépode calanoïde Diaptomus (Aglaodiaptomus)
leptopus S. A. Forbes, 1882. II. Phase copepodite et adulte.
Crustaceana 54:171 – 195.
Plassmann, T., Maier, G., Stich, H. B. 1997. Predation impact of Cyclops vicinus on the rotifer community in Lake Constance in
spring. Journal of Plankton Research 19:1069 – 1079.
Prepas, E. 1978. Sugar-frosted Daphnia: An improved fixation technique for Cladocera. Limnology and Oceanography
23:557 – 559.
Price, H. J., Paffenhöfer, G. -A. 1986. Effects of concentration on the
feeding of a marine copepod in algal monocultures and mixtures. Journal of Plankton Research 8:119 – 128.
Price, H. J., Paffenhöfer, G. -A., Strickler, J. R. 1983. Modes of cell
capture in calanoid copepods. Limnology and Oceanography
28:116 – 123.
Ramalingam, K., Raghunathan, M. B. 1982. A study on the pH tolerance and survival of a cyclopoid copepod Mesocyclops leucarti (Claus). Comparative Physiology and Ecology 7:188 – 190.
Ramcharan, C. W., Sprules, W. G. 1991. Predator-induced behavioral
defense and its ecological consequences for two calanoid copepods. Oecologia 86:276 – 286.
Reddy, Y. R. 1994. Copepoda: Calanoida: Diaptomidae, Key to the
genera Heliodiaptomus, Allodiaptomus, Neodiaptomus, Phyllodiaptomus, Eodiaptomus, Arctodiaptomus and Sinodiaptomus.
Guides to the Identification of the Microinvertebrates of the
Continental Waters of the World. Dumont, H. J. (Ed.) 5:1 – 221.
Reed, E. B. 1963. Records of freshwater Crustacea from Arctic and
Subarctic Canada. National Museum of Canada Bulletin No.
199, Contributions to Zoology 199:29 – 62.
Reed, E. B. 1986. Esteval phenology of an Acanthocyclops (Crustacea, Copepoda) in a Colorado tarn with remarks on the vernalis – robustus complex. Hydrobiologia 139:127 – 133.
Reed, E. B. 1994. Arctodiaptomus novosibiricus Kiefer, 1971 in
Alaska and Northwest Territories, with notes on A. arapahoensis (Dodds, 1915) and a key to New World species of Arctodiaptomus (Copepoda:Calanoida). Proceedings of the Biological
Society of Washington 107:666 – 679.
Reed, E. B. 1995. Cyclops kolensis alaskaensis Lindberg, 1956, revisited (Copepoda: Cyclopoida). Journal of Crustacean Biology
15:365 – 375.
Reed, E. B., Aronson, J. G. 1989. Seasonal variation in length of
copepodids and adults of Diacyclops thomasi (Forbes) in two
Colorado montane reservoirs (Copepoda). Journal of Crustacean Biology 9:67 – 76.
948
C. E. Williamson and J. W. Reid
Reed, E. B., McIntyre, N. E. 1995. Cyclops strenuus (Fischer, 1851)
sensu lato in Alaska and Canada, with new records of occurrence. Canadian Journal of Zoology 73:1699 – 1711.
Reid, J. W. 1986. Some usually overlooked cryptic copepod habitats.
Syllogeus 58:594 – 598.
Reid, J. W. 1988. Copepoda (Crustacea) from a seasonally flooded
marsh in Rock Creek Stream Valley Park, Maryland. Proceedings of the Biological Society of Washington 101:31 – 38.
Reid, J. W. 1989a. The distribution of species of the genus Thermocyclops (Copepoda, Cyclopoida) in the western hemisphere,
with description of T. parvus, new species. Hydrobiologia
175:149 – 174.
Reid, J. W. 1989b. RE:”Infection of a field population of Aedes cantator with a polymorphic microsporidium, Amplyospora connecticus via release of the intermediate copepod host, Acanthocyclops vernalis.” Journal of the American Mosquito Control
Association 5:616 – 617.
Reid, J. W. 1990. Continental and coastal free-living Copepoda
(Crustacea) of Mexico, Central America and the Caribbean region. In: Navarro L. D. Robinson, J. G. (Eds.), Diversidad Biologica en la Reserva de la Biosfera de Sian Ka’an, Quintana
Roo, México. Centro de Investigaciones de Quintana Roo
(CIQRO) and Program of Studies in Tropical Conservation,
University of Florida; Chetumal, Quintana Roo. Pp. 175 – 213.
Reid, J. W. 1991a. The genus Metacyclops (Copepoda:Cyclopoida)
present in North America: M. cushae, new species, from
Louisiana. Journal of Crustacean Biology 11:639 – 646.
Reid, J. W. 1991b. Some species of Tropocyclops (Crustacea, Copepoda) from Brazil, with a key to the American species. Bijdragen tot de Dierkunde 61:3 – 15.
Reid, J. W. 1992. Taxonomic problems: A serious impediment to
groundwater ecological research in North America. In: Stanford
J. A., Simons, J. J. Eds., First International Conference on
Groundwater Ecology. American Water Resources Association,
Bethesda, MD. Pages 133 – 142.
Reid, J. W. 1995. Redescription of Parastenocaris brevipes Kessler
and description of a new species of Parastenocaris (Copepoda:Harpacticoida:Parastenocarididae) from the U.S.A. Canadian Journal of Zoology 73:173 – 187.
Reid, J. W. 1998. How “cosmopolitan” are the continental cyclopoid
copepods? Comparison of North American and Eurasian faunas, with description of Acanthocyclops parasensitivus sp.n.
(Copepoda:Cyclopoida) from the U.S.A. Zoologischer Anzeiger
236(1997):109 – 118.
Reid, J. W., Hare, S. G. F., Nasci, R. S. 1989. Diacyclops navus (Crustacea:Copepoda) redescribed from Louisiana, U.S.A. Transactions of the American Microscopical Society 108:332 – 344.
Reid, J. W., Hunt, G. W., Stanley, E. H. 2001. A new species of Stygonitocrella (Crustacea:Copepoda:Ameiridae) from North
America. Proceedings of the Biological Society of Washington
114 [in press]
Reid, J. W., Ishida, T. 1993. New species and new records of the
genus Elaphoidella (Crustacea:Copepoda:Harpacticoida) from
the United States. Proceedings of the Biological Society of Washington 106:137 – 146.
Reid, J. W., Ishida, T. 1996. Two new species of Gulcamptus (Crustacea:Copepoda:Harpacticoida) from North America. Japanese
Journal of Limnology 57:133 – 144.
Reid, J. W., Ishida, T. 2000. Itocyclops, a new genus proposed for
Speocyclops yezoensis Ito (Copepoda:Cyclopoida). Journal of
Crustacean Biology 20:589 – 596.
Reid, J. W., Janetzky, W. 1996. Colonization of Jamaican bromeliads
by Tropocyclops jamaicensis n. sp. (Crustacea:Copepoda:Cyclopoida). Invertebrate Biology 115:305 – 320.
Reid, J. W., Marten, G. G. 1995. The cyclopoid copepod (Crustacea)
fauna of non-planktonic continental habitats in Louisiana and
Mississippi. Tulane Studies in Zoology and Botany 30:39 – 45.
Reid, J. W., Reed, E. B. 1994. First records of two neotropical species
of Mesocyclops (Copepoda) from Yukon Territory: cases of passive dispersal? Arctic 47:80 – 87.
Reid, J. W., Strayer, D. L., McArthur, J. V., Stibbe, S. 1999. Rheocyclops, a new genus of copepods from the southeastern and central U.S.A. (Copepoda:Cyclopoida). Journal of Crustacean Biology 19:384 – 396.
Richman, S., Dodson, S. I. 1983. The effect of food quality on feeding and respiration by Daphnia and Diaptomus. Limnology and
Oceanography 28:948 – 956.
Riera, T., Estrada, M. 1985. Dimensions and allometry in Tropocyclops prasinus. Empirical relationships with environmental temperature. Verhandlungen der Internationale Vereinigung für
Theoretische und Angewandte Limnologie 22:3159 – 3163.
Ringelberg, J. 1980. Aspects of red pigmentation in zooplankton, especially copepods. In: Kerfoot, W. C. (Ed.), Evolution and ecology of zooplankton communities. Uni. Press of New England,
Hanover, NH. Pages 91 – 97.
Ringelberg, J., Keyser, A. L., Flik, B. J. G. 1984. The mortality effect
of ultraviolet radiation in a translucent and in a red morph of
Acanthodiaptomus denticornis (Crustacea, Copepoda) and its
possible ecological relevance. Hydrobiologia 112:217 – 222.
Robertson, A. 1970. Distribution of calanoid copepods (Calanoida,
Copepoda) in Oklahoma. Proceedings of the Oklahoma Academy of Science 50:98 – 103.
Robertson, A. 1972. Calanoid copepods: new records from Oklahoma. Southwestern Naturalist 17:201 – 203.
Robertson, A. 1975. A new species of Diaptomus (Copepoda,
Calanoida) from Oklahoma and Texas. American Midland Naturalist 93:206 – 214.
Robertson, A., Gannon, J. E. 1981. Annotated checklist of the freeliving copepods of the Great Lakes. Journal of Great Lakes
Research 7:382 – 393.
Rocha, C. E. F. 1991. A new species of Halicyclops (Copepoda,
Cyclopidae) from California, and a revision of some Halicyclops material in the collections of the US Museum of Natural
History. Hydrobiologia 226:29 – 37.
Rocha, C. E. F., Hakenkamp, C. C. 1993. New species of Halicyclops (Copepoda Cyclopidae) from the United States of America. Hydrobiologia 259:145 – 156.
Rocha, C. E. F., Iliffe, T. M., Reid, J. W, Suárez-Morales, E. 1998. A
new species of Halicyclops (Copepoda, Cyclopoida) from
cenotes of the Yucatán Peninsula, Mexico, with an identification key for the species of the genus from the Caribbean region
and adjacent areas. Sarsia 83:387 – 399.
Roche, K. F. 1987. Post-encounter vulnerability of some rotifer prey
types to predation by the copepod Acanthocyclops robustus.
Hydrobiologia 147:229 – 233.
Roff, J. C. 1972. Aspects of the reproductive biology of the planktonic copepod Limnocalanus macrurus Sars, 1863. Crustaceana
22:155 – 160.
Rosenthal, H. 1972. Über die Geschwindigkeit der Sprungbewegungen bei Cyclops strenuus (Copepoda). Internationale Revue der
gesamten Hydrobiologie 57:157 – 167.
Sanders, R. W., Williamson, C. E., Stutzman, P. L., Moeller, R. E.,
Goulden, C. E., Aoki-Goldsmith, R. 1996. Reproductive success
of “herbivorous” zooplankton fed algal and nonalgal food resources. Limnology and Oceanography 41:1295 – 1305.
Santer, B. 1993. Do cyclopoid copepods control Daphnia populations in early spring, thereby protecting their juvenile instar
stages from food limitation? Verhandlungen der Internationale
22. Copepoda
Vereinigung für Theoretische und Angewandte Limnologie
25:634 – 637.
Santer, B. 1994. Influences of food type and concentration on the development of Eudiaptomus gracilis and implications for interactions between calanoid and cyclopoid copepods. Archiv für Hydrobiologie 131:141 – 159.
Santer, B. 1996. Nutritional suitability of the dinoflagellate Ceratium
furcoides for four copepod species. Journal of Plankton Research 18:323 – 333.
Särkkä, J. 1992. Effects of eutrophic and organic loading on the
occurrence of profundal harpacticoids in a lake in southern
Finland. Environmental Monitoring and Assessment 21:
211 – 223.
Sarvala, J. 1979. A parthenogenetic life cycle in a population of Canthocamptus staphylinus (Copepoda, Harpacticoida). Hydrobiologia 62:113 – 129.
Schaffner, W. R., Hairston Jr., N. G., Howarth, R. W. 1994. Feeding
rates and filament clipping by crustacean zooplankton consuming cyanobacteria. Verhandlungen der Internationale Vereinigung für Theoretische und Angewandte Limnologie
25:2375 – 2381.
Schindler, D. W. 1969. Two useful devices for vertical plankton and
water sampling. Journal of the Fisheries Research Board of
Canada 26:1948 – 1955.
Shen, C.-j., Lee, F.-s. 1963. The estuarine Copepoda of Chiekong and
Zaikong Rivers, Kwangtung Province, China. Acta Zoologica
Sinica 15:571 – 596. [In Chinese with English summary].
Shih, C.-T., Maclellan, D. C. 1977. Descriptions of copepodite stages
of Diaptomus (Leptodiaptomus) nudus Marsh 1904 (Crustacea:Copepoda). Canadian Journal of Zoology 55:912 – 921.
Siebeck, O. 1969. Spatial orientation of planktonic crustaceans. I.
The swimming behaviour in a horizontal plane. Verhandlungen
der Internationale Vereinigung für Theoretische und Angewandte Limnologie 17:831 – 840.
Siebeck, O. 1980. Optical orientation of pelagic crustaceans and its
consequences in the pelagic and littoral zones. in: Kerfoot, W.
C. (Ed.), Evolution and ecology of zooplankton communities.
Uni. Press of New England, Hanover, NH. Pages 28 – 38.
Smith, K. E., Fernando, C. H. 1978. A guide to the freshwater
calanoid and cyclopoid Copepoda Crustacea of Ontario. University of Waterloo Biology Series 18:1 – 76.
Smyly, W. J. P. 1962. Laboratory experiments with stage V copepodites
of the freshwater copepod, Cyclops leuckarti Claus, from Windermere and Esthwaite Water. Crustaceana 4:273 – 280.
Smyly, W. J. P. 1980. Effect of constant and alternating temperatures
on adult longevity of the freshwater cyclopoid copepod, Acanthocyclops viridis (Jurine). Archiv für Hydrobiologie
89:353 – 362.
Soto, D., Hurlbert, S. H. 1991. Long-term experiments on calanoidcyclopoid interactions. Ecological Monographs 61:245 – 265.
Stearns, D. E., Sharp, A. A. 1994. Sublethal effects of cupric ion activity on the phototaxis of three calanoid copepods. Hydrobiologia 292 / 293:505 – 511.
Steib, K., Mayer, P. 1988. Epidemiology and vectors of Dracunculus
medinensis in northwest Burkina Faso, West Africa. Annals of
Tropical Medicine and Parasitology 82:189 – 199.
Stemberger, R. S. 1985. Prey selection by the copepod Diacyclops
thomasi. Oecologia 65:492 – 497.
Stemberger, R. S. 1986. The effects of food deprivation, prey density
and volume on clearance rates and ingestion rates of Diacyclops
thomasi. Journal of Plankton Research 8:243 – 251.
Stemberger, R. S. 1995. Pleistocene refuge areas and postglacial dispersal of copepods of the northeastern United States. Canadian
Journal of Fisheries and Aquatic Sciences 52:2197 – 2210.
949
Stemberger, R. S., Gilbert, J. J. 1984. Spine development in the rotifer
Keratella cochlearis: induction by cyclopoid copepods and Asplanchna. Freshwater Biology 14:639 – 647.
Stemberger, R. S., Lazorchak, J. M. 1994. Zooplankton assemblage
responses to disturbance gradients. Canadian Journal of Fisheries and Aquatic Sciences 51:2435 – 2447.
Sterner, R. W., Hessen, D. O. 1994. Algal nutrient limitation and the
nutrition of aquatic herbivores. Annual Review of Ecology and
Systematics 25:1 – 29.
Strayer, D. L. 1985. The benthic micrometazoans of Mirror Lake, New
Hampshire. Archiv für Hydrobiologie, Supplement 72:287 – 426.
Strayer, D. L., Reid, J. W. 1999. Distribution of hyporheic cyclopoids
(Crustacea: Copepoda) in the eastern United States. Archiv für
Hydrobiologie 145:79 – 92.
Strickler, J. R. 1975a. Intra- and interspecific information flow
among planktonic copepods: Receptors. Verhandlungen der Internationale Vereinigung für Theoretische und Angewandte
Limnologie 19:2951 – 2958.
Strickler, J. R. 1975b. Swimming of planktonic Cyclops species (Copepoda, Crustacea): Pattern, movements and their control. In: Wu,
T. Y., Brokaw, C. J., Brennan, C. (Eds.), Swimming and flying in
nature. Vol. 2. Plenum Press, New York. Pages 599 – 613.
Strickler, J. R. 1982. Calanoid copepods, feeding currents, and the
role of gravity. Science 218:158 – 160.
Strickler, J. R., Bal, A. K. 1973. Setae of the first antennae of the
copepod Cyclops scutifer (Sars): Their structure and importance. Proceedings of the National Academy of Science of the
U.S.A. 70:2656 – 2659.
Stutzman, P. 1995. Food quality of gelatinous colonial chlorophytes
to the freshwater zooplankters Daphnia pulicaria and Diaptomus oregonensis. Freshwater Biology 34:149 – 153.
Suárez, M. F., Marten, G. G., Clark, G. G. 1992. A simple method
for cultivating freshwater copepods used in biological control of
Aedes aegypti. Journal of the American Mosquito Control Association 8:409 – 412.
Suárez-Morales, E., Elías-Gutiérrez, M. 2000. Two new Mastigodiaptomus (Copepoda, Diaptomidae) from southeastern Mexico,
with a key for the identification of the know on species of the
genus. Journal of Natural History 34:693 – 708.
Suárez-Morales, E., Reid, J. W. 1998. An updated list of the free-living freshwater copepods (Crustacea) of Mexico. Southwestern
Naturalist 43:256 – 265.
Suárez-Morales, E., Reid, J. W., Iliffe, T. M., Fiers, F. 1996. Catálogo
de los Copépodos (Crustacea) Continentales de la Península de
Yucatán, México. Comisión Nacional para el Conocimiento y
Uso de la Biodiversidad (CONABIO) and El Colegio de la Frontera Sur (ECOSUR), Mexico City. 296 p.
Svensson, J.-E. 1997. Chaoborus predation and sex-specific mortality
in a copepod. Limnology and Oceanography 42:572 – 577.
Tessier, A. J., Goulden, C. E. 1982. Estimating food limitation in cladoceran populations. Limnology and Oceanography 27:707 – 717.
Threlkeld, S. T., Dirnberger, J. M. 1986. Benthic distributions of
planktonic copepods, especially Mesocyclops edax. Syllogeus
58:481 – 486.
Tinson, S., Laybourn-Parry, J. 1985. The behavioural responses and
tolerance of freshwater benthic cyclopoid copepods to hypoxia
and anoxia. Hydrobiologia 127:257 – 263.
Torke, B. G. 1976. A key to the identification of the cyclopoid copepods of Wisconsin, with notes on their distribution and ecology.
Wisconsin Department of Natural Resources Research Report
88:1 – 15.
Twombly, S., Burns, C. W. 1996. Exuvium analysis: A nondestructive
method of analyzing copepod growth and development. Limnology and Oceanography 41:1324 – 1329.
950
C. E. Williamson and J. W. Reid
Vanderploeg, H. A. 1994. Zooplankton particle selection and feeding
mechanisms. In R. S. Wotton (Ed.), The biology of particles in
aquatic systems. Lewis, Ann Arbor, MI. Pages 205 – 234.
Vanderploeg, H. A., Bolsenga, S. J., Fahnenstiel, G. L., Liebig, J. R.,
Gardner, W. S. 1992a. Plankton ecology in an ice-covered bay
of Lake Michigan: utilization of a winter phytoplankton bloom
by reproducing copepods. Hydrobiologia 243 / 244:175 – 183.
Vanderploeg, H. A., Gardner, W. S., Parrish, C. C., Liebig, J. R., Cavaletto, J. F. 1992b. Lipids and life-cycle strategy of a hypolimnetic copepod in Lake Michigan. Limnology and Oceanography
37:413 – 424.
Vanderploeg, H. A., Liebig, J. R., Omair, M. 1993. Bythotrephes predation on Great Lakes’ zooplankton measured by an in situ
method: implications for zooplankton community structure.
Archiv für Hydrobiologie 127:1 – 8.
Vanderploeg, H. A., Paffenhöfer, G.-A. 1985. Modes of algal capture
by the freshwater copepod Diaptomus sicilis and their relation
to food-size selection. Limnology and Oceanography 30:
871 – 885.
Vanderploeg, H. A., Paffenhöfer, G.-A., Liebig, J. R. 1988. Diaptomus vs net phytoplankton: effects of algal size and morphology
on selectivity of a behaviorally flexible, omnivorous copepod.
Bulletin of Marine Science 43:377 – 394.
Vanderploeg, H. A., Scavia, D., Liebig, J. R. 1984. Feeding rate of
Diaptomus sicilis and its relation to selectivity and effective
food concentration in algal mixtures and in Lake Michigan.
Journal of Plankton Research 6:919 – 941.
van Leeuwen, H. C., Maly, E. J. 1991. Changes in swimming behavior of male Diaptomus leptopus (Copepoda:Calanoida) in response to gravid females. Limnology and Oceanography
36:1188 – 1195.
Vitousek, P. M. 1994. Beyond global warming: ecology and global
change. Ecology 75:1861 – 1876.
Vu, S. N., Nguyen, T. Y., Kay, B. H., Marten, G. G., Reid, J. W.
1998. Eradication of Aedes aegypti from a village in Vietnam,
using copepods and community participation. American Journal
of Tropical Medicine and Hygiene 59:657 – 660.
Walter, T. C. 1989. Review of the New World species of Pseudodiaptomus (Copepoda: Calanoida), with a key to the species. Bulletin of Marine Science 45:590 – 628.
Walton, W. E. 1985. Factors regulating the reproductive phenology
of Onychodiaptomus birgei (Copepoda: Calanoida). Limnology
and Oceanography 30:167 – 179.
Watras, C. J. 1983a. Mate location by diaptomid copepods. Journal
of Plankton Research 5:417 – 423.
Watras, C. J. 1983b. Reproductive cycles in diaptomid copepods:
Effects of temperature, photocycle, and species on reproductive
potential. Canadian Journal of Fisheries and Aquatic Sciences
40:1607 – 1613.
Watras, C. J., Haney, J. F. 1980. Oscillations in the reproductive
condition of Diaptomus leptopus (Copepoda: Calanoida) and
their relation to rates of egg-clutch production. Oecologia
45:94 – 103.
Wetzel, R. G. 1983. Limnology, 2nd ed. Saunders, Philadelphia, PA.
767 p.
Whitehouse, J. W., Lewis, B. G. 1973. The effect of diet and density
on development, size and egg production in Cyclops abyssorum
Sars, 1863 (Copepoda, Cyclopoida). Crustaceana 25:225 – 236.
Wickham, S. A. 1995. Cyclops predation on ciliates: species-specific
differences and functional responses. Journal of Plankton Research 17:1633 – 1646.
Wierzbicka, M., 1962. On the resting stage and mode of life of some
species of Cyclopoida. Polish Archives of Hydrobiology (Polskie
Archiwum Hydrobiologii) 10:215 – 229.
Williamson, C. E. 1980. The predatory behavior of Mesocyclops
edax: Predator preferences, prey defenses, and starvationinduced changes. Limnology and Oceanography 25:903 – 909.
Williamson, C. E. 1981. Foraging behavior of a freshwater copepod:
Frequency changes in looping behavior at high and low prey
densities. Oecologia 50:332 – 336.
Williamson, C. E. 1983a. Behavioural interactions between a cyclopoid copepod predator and its prey. Journal of Plankton
Research 5:701 – 711.
Williamson, C. E. 1983b. Invertebrate predation on planktonic rotifers. Hydrobiologia 104:385 – 396.
Williamson, C. E. 1984. Laboratory and field experiments on the
feeding ecology of the freshwater cyclopoid copepod, Mesocyclops edax. Freshwater Biology 14:575 – 585.
Williamson, C. E. 1986. The swimming and feeding behavior of
Mesocyclops. Hydrobiologia 134:11 – 19.
Williamson, C. E. 1987. Predator – prey interactions between omnivorous diaptomid copepods and rotifers: The role of prey
morphology and behavior. Limnology and Oceanography
32:167 – 177.
Williamson, C. E. 1993. Linking predation risk models with behavioral mechanisms: identifying population bottlenecks. Ecology
74:320 – 331.
Williamson, C. E., Butler, N. M. 1986. Predation on rotifers by the
suspension-feeding calanoid copepod Diaptomus pallidus. Limnology and Oceanography 31:393 – 402.
Williamson, C. E., Butler, N. M. 1987. Temperature, food, and mate
limitation of copepod reproductive rates: separating the effects
of multiple hypotheses. Journal of Plankton Research 9:
821 – 836.
Williamson, C. E., Butler, N. M., Forcina, L. 1985. Food limitation
in naupliar and adult Diaptomus pallidus. Limnology and
Oceanography 30:1283 – 1290.
Williamson, C. E., Magnien, R. E. 1982. Diel vertical migration in
Mesocyclops edax: Implications for predation rate estimates.
Journal of Plankton Research 4:329 – 339.
Williamson, C. E., Stemberger, R. S., Morris, D. P., Frost, T. M.,
Paulsen, S. G. 1996. Ultraviolet radiation in North American
lakes: attenuation estimates from DOC measurements and implications for plankton communities. Limnology and Oceanography 41:1024 – 1034.
Williamson, C. E., Stoeckel, M. E., Schoeneck, L. J. 1989. Predation
risk and the structure of freshwater zooplankton communities.
Oecologia 79:76 – 82.
Williamson, C. E., Vanderploeg, H. A. 1988. Predatory suspensionfeeding in Diaptomus: prey defenses and the avoidance of cannibalism. Bulletin of Marine Science 43:561 – 572.
Williamson, C. E., Zagarese, H. E., Schulze, P. C., Hargreaves, B. R.,
Seva, J. 1994. The impact of short-term exposure to UV-B radiation on zooplankton communities in north temperate lakes.
Journal of Plankton Research 16:205 – 218.
Wilson, C. B. 1932. The copepods of the Woods Hole region, Massachusetts. Bulletin of the United States National Museum
158:1 – 635 and plates 1 – 41.
Wilson, C. C., Hebert, P. D. N. 1993. Impact of copepod predation
on distribution patterns of Daphnia pulex clones. Limnology
and Oceanography 38:1304 – 1310.
Wilson, M. S. 1958. The copepod genus Halicyclops in North America, with description of a new species from Lake Pontchartrain,
Louisiana, and the Texas coast. Tulane Studies in Zoology
6:176 – 189.
Wilson, M. S., Yeatman, H. C. 1959. Free-living Copepoda. In Edmondson, W. T. (Ed.), Ward and Whipple’s Fresh-water biology.
J Wiley, New York, Pages 735 – 868.
22. Copepoda
Winfield, I. J., Townsend, C. R. 1983. The cost of copepod reproduction: increased susceptibility to fish predation. Oecologia
60:406 – 411.
Wong, C. K. 1981. Predatory feeding behavior of Epischura lacustris
(Copepoda, Calanoida) and prey defense. Canadian Journal of
Fisheries and Aquatic Sciences 38:275 – 279.
Wong, C. K., Sprules, W. G. 1986. The swimming behavior of the
freshwater calanoid copepods Limnocalanus macrurus Sars,
Senecella calanoides Juday and Epischura lacustris Forbes. Journal of Plankton Research 8:79 – 90.
Woodmansee, R. A., Grantham, B. J. 1961. Diel vertical migrations
of two zooplankters (Mesocyclops and Chaoborus) in a Mississippi lake. Ecology 42:619 – 628.
Woodward, I. O., White, R. W. G. 1981. Effects of temperature and
food on the fecundity and egg development rates of Boeckella
symmetrica Sars (Copepoda: Calanoida). Australian Journal of
Marine and Freshwater Research 32:997 – 1002.
Wright, D., O’Brien, W. J., Vinyard, G. L. 1980. Adaptive value of
vertical migration: A simulation model argument for the predation hypothesis. In: W. C. Kerfoot (Ed.), Evolution and ecology
of zooplankton communities. Uni. Press of New England,
Hanover, NH. Pages 138 – 147.
Wyngaard, G. A. 1983. In situ life table of a subtropical copepod.
Freshwater Biology 13:275 – 281.
Wyngaard, G. A. 1986. Genetic differentiation of life history traits in
TABLE I
951
populations of Mesocyclops edax (Crustacea: Copepoda). Biological Bulletin 170:279 – 295.
Wyngaard, G. A. 1998. Reciprocal transfer study of north temperate
and subtropical populations of Mesocyclops edax (Copepoda:
Cyclopoida). Journal of Marine Systems 15:163 – 169.
Wyngaard, G. A., Chinnappa, C. C. 1982. General biology and cytology of cyclopoids. In: Harrison, F. W., Cowden, R. R. (Eds.),
Developmental biology of freshwater invertebrates. A. R. Liss,
New York. Pages 485 – 533.
Wyngaard, G. A., Taylor, B. E., Mahoney, D. L. 1991. Emergence
and dynamics of cyclopoid copepods in an unpredictable environment. Freshwater Biology 25:219 – 232.
Yan, N. D., Mackie, G. L., Dillon, P. J. 1990. Cadmium concentrations of crustacean zooplankton of acidified and nonacidified
Canadian Shield lakes. Environmental Science and Technology
24:1367 – 1372.
Yan, N. D., Pawson, T. W. 1997. Changes in the crustacean zooplankton community of Harp Lake, Canada, following invasion
by Bythotrephes cederstroemi. Freshwater Biology 37:409 – 425.
Zaret, T. M. 1980. Predation in freshwater communities. Yale Uni.
Press, New Haven, CT, 187 pp.
Zaret, T. M., Suffern, J. S. 1976. Vertical migration in zooplankton
as a predator avoidance mechanism. Limnology and Oceanography 21:804 – 813.
Orders and Superorders of Copepods
Platycopioida Fosshagen 1985 — Small, hyperbenthic marine species in shallow seas and anchialine caves.
Calanoida Sars 1903 — Freshwater and marine, free-swimming.
Misophrioida Gurney 1933 — shallow coastal waters, deep sea, deep-water plankton, and anchialine caves.
Harpacticoida Sars 1903 — Freshwater and marine, free-living, primarily benthic or attached.
Monstrilloida Sars 1903 — nauplii and copepodids are endoparasitic on polychaetes and prosobranch molluscs. Adults are free-swimming, but
nonfeeding. Adult females carry eggs on spines.
Mormonilloida Boxshall 1979 — only two species known, both widely distributed and abundant in deep oceans (400 – 1500 m depth). They are
free-swimming particle-feeders.
Gelyelloida Huys 1988 — only three species known, two from Europe (France and Switzerland) and one from the United States (South Carolina). All are freshwater and benthic / subterranean.
Cyclopoida Burmeister 1835 — Freshwater and marine, benthic and planktonic.
Siphonostomatoida Thorell 1859 — parasitic or associated with fish or invertebrate hosts; primarily marine, but some freshwater species exist.
Body often highly reduced, lacking appendages.
Poecilostomatoida Thorell 1859 — Mostly parasitic on, or associated with, a wide variety of fish and invertebrates. Morphologically the most
diverse copepod order; mostly marine. Four families are often abundant in the marine plankton. Some feed visually (Corycaeidae and Sapphirinidae). Includes the family Ergasilidae.
Superorder Gymnoplea — Articulation between prosome and urosone is between fifth pedigerous and genital somites. This is the ancestral tagmosis that includes the Platycopioida and Calanoida.
Superorder Podoplea — Articulation between prosome and urosome is between fourth and fifth pedigerous somites. This is the derived tagmosis,
and it includes the other eight orders.
From Huys and Boxshall, 1991.
952
C. E. Williamson and J. W. Reid
TABLE II
North American Copepods of the Order Calanoida G. O. Sars, 1903
Family Acartiidae G. O. Sars, 1903
Acartia: sinensis Shen and Lee, 1963 (I)
Family Aetideidae Giesbrecht, 1892
Senecella: calanoides Juday, 1923
Family Centropagidae Giesbrecht, 1892
Limnocalanus: johanseni Marsh, 1920; macrurus G. O. Sars, 1863.
Osphranticum: labronectum S. A. Forbes, 1882.
Sinocalanus: doerri (Brehm, 1909) (I) (see Orsi et al., 1983).
Family Diaptomidae G. O. Sars, 1903
Acanthodiaptomus: denticornis (Wierzejski, 1887).
Aglaodiaptomus: atomicus DeBiase and Taylor, 1997; clavipes (Schacht, 1897); clavipoides M. S. Wilson, 1955; conipedatus (Marsh, 1907);
dilobatus M. S. Wilson, 1958; forbesi Light, 1938; kingsburyae Robertson, 1975; leptopus (S. A. Forbes, 1882); lintoni (S. A. Forbes,
1893); marshianus M. S. Wilson, 1953; pseudosanguineus (Turner, 1921); saskatchewanensis M. S. Wilson, 1958; savagei DeBiase and
Taylor, 2000; spatulocrenatus (Pearse, 1906); stagnalis (S. A. Forbes, 1882).
Arctodiaptomus: arapahoensis (Dodds, 1915); bacillifer (Koelbel, 1884); dorsalis (Marsh, 1907); floridanus (Marsh, 1926); kurilensis Kiefer,
1937; saltillinus (Brewer, 1898). Key: Reddy, 1994, Reed, 1994.
Diaptomus: glacialis Lilljeborg, 1889.
Eudiaptomus: gracilis (G. O. Sars, 1863); yukonensis Reed, 1991.
Hesperodiaptomus: arcticus (Marsh, 1920); augustaensis (Turner, 1910); breweri M. S. Wilson, 1958; caducus Light, 1938; californiensis
Scanlin and Reid, 1996; eiseni (Lilljeborg, 1889); franciscanus (Lilljeborg, 1889); hirsutus M. S. Wilson, 1953; kenai M. S. Wilson, 1953;
kiseri (Kincaid, 1953); nevadensis Light 1938; novemdecimus M. S. Wilson, 1953; schefferi M. S. Wilson, 1953; shoshone (S. A. Forbes,
1893); victoriaensis Reed, 1958; wardi (Pearse, 1905); wilsonae Reed, 1958.
Leptodiaptomus: ashlandi (Marsh, 1893); assiniboiaensis (Anderson, 1970); coloradensis (Marsh, 1911); connexus Light, 1938; cuauhtemoci
(Osorio-Tafall, 1941; synonym assiniboiaensis Anderson, 1970); insularis Kincaid, 1956; judayi (Marsh, 1907); minutus (Lilljeborg, 1889);
moorei M. S. Wilson, 1954; novamexicanus (Herrick, 1895); nudus (Marsh, 1904); pribilofensis (Juday and Muttkowski, 1915); sicilis (S.
A. Forbes, 1882); siciloides (Lilljeborg, 1889); signicauda (Lilljeborg, 1889); spinicornis Light, 1938; trybomi (Lilljeborg, 1889); tyrrelli
(Poppe, 1888).
Mastigodiaptomus: albuquerquensis (Herrick, 1895); texensis M. S. Wilson, 1953.Key: Suárez-Morales and Elías-Gutiérrez, 2000.
Mixodiaptomus: theeli (Lilljeborg, 1889).
Nordodiaptomus: alaskaensis M. S. Wilson, 1951.
Onychodiaptomus: birgei (Marsh, 1894); hesperus M. S. Wilson and Light, 1951; louisianensis M. S. Wilson and Moore, 1953; sanguineus
(S. A. Forbes, 1876); virginiensis (Marsh, 1915).
Sinodiaptomus: sarsi (Rylov, 1923)(I). Key: Reddy, 1994.
Skistodiaptomus: bogalusensis M. S. Wilson and Moore, 1953; bogalusensis marii Harris, 1978; carolinensis Yeatman, 1986; mississippiensis
(Marsh, 1894); oregonensis (Lilljeborg, 1889); pallidus (Herrick, 1879); pygmaeus (Pearse, 1906); reighardi (Marsh, 1895); sinuatus Kincaid, 1953.
Family Temoridae Giesbrecht, 1892
Epischura: fluviatilis Herrick, 1883; lacustris S. A. Forbes, 1882; massachusettsensis Pearse, 1906; nevadensis Lilljeborg, 1889; nordenskioldi
Lilljeborg, 1889.
Eurytemora: affinis (Poppe, 1880); americana Williams, 1906; arctica M. S. Wilson and Tash, 1966; bilobata Akatova, 1949 (synonym yukonensis M. S. Wilson, 1953); canadensis Marsh, 1920; composita Keiser, 1929; gracilicauda Akatova, 1949.
Heterocope: septentrionalis Juday and Muttkowski, 1915.
Identification is based primarily on males. Species primarily occurring in brackish coastal waters, some of which may occasionally enter fresh
water, are not included. Introduced species are indicated by (I). Keys and taxonomic revisions published after 1959 are noted.
22. Copepoda
TABLE III
953
North American Copepods of the Order Cyclopoida Burmeister, 1834
Family Cyclopidae Dana, 1853
Acanthocyclops: brevispinosus (Herrick, 1884); capillatus (G. O. Sars, 1863); carolinianus Yeatman, 1944; columbiensis Reid, 1990; exilis
(Coker, 1934); montana Reid and Reed in Reid et al., 1991; parasensitivus Reid, 1998; parvulus Strayer, 1989; pennaki Reid, 1992;
robustus (G. O. Sars, 1863); venustoides (Coker, 1934); vernalis (Fischer, 1853). Key: Einsle, 1996a.
Apocyclops: dimorphus (Kiefer, 1931); panamensis (Marsh, 1913); spartinus Ruber, 1968.
Bryocyclops: muscicola (Menzel, 1926). (I)
Cryptocyclops: bicolor (G. O. Sars, 1862).
Cyclops: canadensis Einsle, 1988; columbianus Lindberg, 1956 (inadequately described species); furcifer (Claus, 1857); kolensis alaskaensis
Lindberg, 1956; laurenticus Lindberg, 1956 (inadequately described); scutifer G. O. Sars, 1863; strenuus Fischer, 1851; vicinus Uljanin,
1875. Revisions: Reed (1995), Reed and McIntyre (1995); Key: Einsle, 1996a.
Diacyclops: alabamensis Reid, 1992; albus Reid, 1992; bernardi (Petkovski, 1986); bicuspidatus (Claus, 1857); bicuspidatus lubbocki (Brady,
1868); bisetosus (Rehberg, 1880); chrisae Reid, 1992; crassicaudis (G. O. Sars, 1863); crassicaudis var. brachycercus (Kiefer, 1927);
dimorphus Reid and Strayer, 1994; harryi Reid, 1992; haueri (Kiefer, 1931); hypnicola (Gurney, 1927); jeanneli (Chappuis, 1929); jeanneli
putei (Yeatman, 1943); languidoides (Lilljeborg, 1901) s.l.; languidus (G. O. Sars, 1863); nanus (G. O. Sars, 1863); navus (Herrick, 1882);
nearcticus Kiefer, 1934; palustris Reid, 1988; sororum Reid, 1992; thomasi (S. A. Forbes, 1882); yeatmani Reid, 1988; Key: Reid, 1988,
emend. Reid et al., 1989.
Ectocyclops: phaleratus (Koch, 1838); polyspinosus Harada, 1931; rubescens Brady, 1904.
Eucyclops: agilis (Koch, 1838); agilis montanus (Brady, 1878); bondi Kiefer, 1934; conrowae Reid, 1992; elegans (Herrick, 1884; synonym E.
neomacruroides Dussart and Fernando, 1990); macruroides denticulatus (Graeter, 1903); prionophorus Kiefer, 1931.
Homocyclops: ater (Herrick, 1882).
Itocyclops: yezoensis (Ito, 1954) (see Reid and Ishida, 2000).
Macrocyclops: albidus (Jurine, 1820); fuscus (Jurine, 1820).
Megacyclops: donnaldsoni (Chappuis, 1929); gigas (Claus, 1857); latipes (Lowndes, 1927); magnus (Marsh, 1920); viridis (Jurine, 1820)(I).
Key: Einsle, 1996a.
Mesocyclops: americanus Dussart, 1985; edax (S. A. Forbes, 1891); longisetus (Thiébaud, 1912); longisetus var. curvatus Dussart, 1987; ruttneri Kiefer, 1981 (I); venezolanus Dussart, 1987. Key: Reid and Reed, 1994.
Metacyclops: cushae Reid, 1991; in Reid, 1991a.
Microcyclops: pumilis Pennak and Ward, 1985; rubellus (Lilljeborg, 1901); varicans (G. O. Sars, 1862).
Orthocyclops: modestus (Herrick, 1883).
Paracyclops: canadensis (Willey, 1934); chiltoni (Thomson, 1882); fimbriatus (Fischer, 1853); poppei (Rehberg, 1880); smileyi Strayer, 1989;
yeatmani Daggett and Davis, 1974. Key: Karaytug, 1999.
Rheocyclops: carolinianus Reid, in Reid et al., 1999; hatchiensis Reid and Strayer in Reid et al., 1999; talladega Reid and Strayer in Reid
et al., 1999; virginianus (Reid, 1993). Key: Reid et al., 1999.
Stolonicyclops: heggiensis Reid and Spooner, 1998.
Thermocyclops: crassus (Fischer, 1853) (I); inversus Kiefer, 1936; parvus Reid, 1989; tenuis (Marsh, 1909). Revision: Reid (1989a); Key:
Herbst, 1986.
Tropocyclops: extensus Kiefer, 1931; extensus f. longispina Kiefer, 1931; jerseyensis Kiefer, 1931; prasinus (Fischer, 1860); prasinus
mexicanus Kiefer, 1938. Key: Reid, 1991b.
Family Oithonidae Dana, 1853
Limnoithona: sinensis (Burckhardt, 1912) (I).
Identification is based primarily on females. Species primarily occurring in brackish coastal waters, some of which may occasionally enter freshwater, are not included. Introduced species are indicated by (I). Keys and taxonomic revisions published after 1959 are noted.
954
C. E. Williamson and J. W. Reid
TABLE IV
North American Copepods of the Order Harpacticoida G. O. Sars, 1903
Family Ameiridae Monard, 1927
Nitocrellopsis: texana Fiers and lliffe, 2000
Nitokra: hibernica (Brady, 1880) (I); lacustris (Shmankevich, 1875).
Psammonitocrella: boultoni Rouch, 1992; longifurcata Rouch, 1992.
Stygonitocrella sequoyahi Reid, Hunt, and Stanley, 2001.
Family Canthocamptidae Brady, 1880
Attheyella: americana (Herrick, 1884); carolinensis Chappuis, 1932; dentata (Poggenpol, 1874); dogieli (Rylov, 1923); idahoensis (Marsh,
1903); illinoisensis (S. A. Forbes, 1882); nordenskioldi (Lilljeborg, 1902); obatogamensis (Willey, 1925); pilosa Chappuis, 1929; spinipes
Reid, 1987; trispinosa (Brady, 1880); ussuriensis Rylov, 1933.
Bryocamptus: arcticus (Lilljeborg, 1902); calvus (Brehm, 1927); cuspidatus (Schmeil, 1893); douwei (Willey, 1925); hiatus (Willey, 1925);
hiemalis (Pearse, 1905); hutchinsoni Kiefer, 1929; minnesotensis (Herrick, 1884; inadequately described species); minusculus (Willey,
1925); minutus (Claus, 1863); morrisoni (Chappuis, 1929); morrisoni elegans (Chappuis, 1929); newyorkensis (Chappuis, 1927); nivalis
(Willey, 1925); pilosus Flössner, 1989; pygmaeus (G. O. Sars, 1862); subarcticus (Willey, 1925); tikchikensis M. S. Wilson, 1958; umiatensis M. S. Wilson, 1958; vejdovskyi (Mrázek, 1893); vejdovskyi f. minutiformis Kiefer, 1934; washingtonensis M. S. Wilson, 1958;
zschokkei (Schmeil, 1893); zschokkei alleganiensis Coker, 1934.
Canthocamptus: assimilis Kiefer, 1931; oregonensis M. S. Wilson, 1956; robertcokeri M. S. Wilson, 1958; sinuus Coker, 1934; staphylinoides
Pearse, 1905; staphylinus (Jurine, 1820) (records of staphylinus in North America are doubtful); vagus Coker and Morgan, 1940.
Elaphoidella: amabilis Ishida in: Reid and Ishida, 1993; bidens (Schmeil, 1894); californica M. S. Wilson, 1975; carterae Reid in: Reid and
Ishida, 1993; fluviusherbae Bruno and Reid, in Bruno et al., 2000); kodiakensis M. S. Wilson, 1975; marjoryae Bruno and Reid, in Bruno
et al., 2000); reedi M. S. Wilson, 1975; shawangunkensis Strayer, 1989; subgracilis (Willey, 1934); tenuicaudis (Herrick, 1884; inadequately described species); wilsonae Hunt, 1979. Key: Reid and Ishida, 1993.
Epactophanes: richardi Mrázek, 1893.
Gulcamptus: alaskaensis Ishida, 1996 in: Reid and Ishida, 1996; huronensis Reid, 1996 in Reid and Ishida, 1996; laurentiacus (Flössner,
1992). Revision: Reid and Ishida, 1996.
Maraenobiotus: brucei (Richard, 1898); canadensis Flössner, 1992; insignipes (Lilljeborg, 1902).
Mesochra: alaskana M. S. Wilson, 1958.
Moraria: affinis Chappuis, 1927; arctica Flössner, 1989; cristata Chappuis, 1929; duthiei (T. and A. Scott, 1896); laurentica Willey, 1927;
mrazeki T. Scott, 1902; virginiana Carter, 1944.
Paracamptus: reductus M. S. Wilson, 1956; reggiae M. S. Wilson, 1958.
Family Canthocamptidae incertae sedis:
Cletocamptus: albuquerquensis (Herrick, 1895); brevicaudata (Herrick, 1895); deitersi (Richard, 1897).
Family Huntemanniidae Por, 1986
Huntemannia: lacustris M. S. Wilson, 1958.
Nannopus: palustris Brady, 1880.
Family Laophontidae T. Scott, 1904
Onychocamptus: mohammed (Blanchard and Richard, 1891).
Family Parastenocarididae Chappuis, 1940
Parastenocaris: brevipes Kessler, 1913; delamarei Chappuis in Chappuis and Delamare Deboutteville, 1958; lacustris Chappuis in Chappuis
and Delamare Deboutteville, 1958; palmerae Reid, 1992; texana Whitman, 1984; trichelata Reid, 1995.
Family Phyllognathopodidae Gurney, 1932
Phyllognathopus: viguieri (Maupas, 1892).
Identification is based on either sex. Species primarily occurring in brackish coastal waters, some of which may occasionally enter freshwater,
are not included. Introduced species are indicated by (I). Keys and taxonomic revisions published after 1959 are noted.
TABLE V
North American Copepods of the Order Poecilostomatoida Thorell, 1859
Family Ergasilidae Nordmann, 1832
Ergasilus: arthrosis Roberts, 1969; auritus Markevich, 1940; caeruleus C. B. Wilson, 1911; celestis Mueller, 1937; centrarchidarum Wright,
1882; cerastes Roberts, 1969; chautauquaensis Fellows, 1887; clupeidarum Johnson and Rogers, 1972; cotti Kellicott, 1892; cyprinaceus
Rogers, 1969; elongatus C. B. Wilson, 1916; japonicus Harada, 1930 (I); lanceolatus C. B. Wilson, 1916; luciopercarum Henderson, 1926;
megaceros C. B. Wilson, 1916; nerkae Roberts, 1963; rhinos Burris and Miller, 1972; sieboldi Nordmann, 1832; tenax Roberts, 1965;
versicolor C. B. Wilson, 1911; wareaglei Johnson, 1971.
Adult females are ectoparasites of fishes, but males and copepodids are planktonic. Identification is based on females. Species primarily occurring in brackish coastal waters, some of which may occasionally enter freshwater, are not included. Introduced species are indicated by (I). No
keys or taxonomic revisions have been published since 1959.
23
DECAPODA
H. H. Hobbs III
Department of Biology
Wittenberg University
Springfield, Ohio 45501
I. Introduction
II. Anatomy and Physiology
A. External Morphology
B. Organ System Function
C. Aspects of Decapod Physiology
III. Ecology and Evolution
A. Diversity, Distribution, and
Evolutionary Relationships
B. Reproduction and Life History
C. Ecological Interactions
I. INTRODUCTION
The Decapoda (Latreille) represent a very significant order that is assigned to the class Malacostraca
and encompasses an immense diversity of marine,
freshwater, and semiterrestrial crustaceans, with some
10,000 species having been described. The two infraorders treated in this chapter, Caridea and Astacidea,
are nearly worldwide in distribution and for purposes
of this discussion are represented by freshwater
shrimps and crayfishes (see Bowman and Abele, 1982
for a detailed classification; see also Hart, 1994). These
aquatic arthropods have successfully invaded a wide
variety of aquatic and semiaquatic habitats, occurring
as obligate cave-dwellers [stygobites ( stygobionts)],
as stream, lake, pond, and swamp dwellers, and as primary burrowers; a few even invade saline environments. The crayfishes and one genus of shrimps (Macrobrachium), in particular, attain the greatest size
among the freshwater crustaceans in North America.
Summarized in this chapter are the ecology and distribution of shrimps and crayfishes in North American
freshwaters (treatment generally restricted to the United
States). Species richness and niche diversification are
particularly well demonstrated in aquatic ecosystems of
Ecology and Classification of North American Freshwater Invertebrates. 2nd Edition
Copyright © 2001 by Academic Press. All rights of reproduction in any form reserved.
IV. Current and Future Research Problems
V. Collecting and Rearing Techniques
A. Shrimps
B. Crayfishes: Astacidae
and Cambaridae
VI. Identification
A. Preparation of Specimens
B. Taxonomic Key to Genera
of Freshwater Decapoda
Literature Cited
the southeastern United States. That these crustaceans
have successfully colonized such diverse habitats is reflected in various morphological and physiological
adaptations discussed. Life-history patterns are reviewed, particularly with reference to how they are influenced by environmental conditions. An examination
is made of the role of these decapods in the functioning
of various aquatic communities. As might be anticipated, a much larger body of data is available for crayfishes than for shrimps and is reflected in the treatment
of these groups. Under each major section of the chapter, a discussion of shrimps appears first, followed by
that of crayfishes. Where data are somewhat limited
(particularly with reference to shrimps), both groups are
discussed in the same section.
II. ANATOMY AND PHYSIOLOGY
A. External Morphology
Although decapods represent the greatest diversity
among orders of crustaceans, all possess a number of
common characteristics, including a carapace that encloses the bronchial chamber. In addition, the first three
955
956
H. H. Hobbs III
pairs of thoracic appendages are modified in all species
as maxillipeds. Carideans (Fig. 1) are easily distinguished from other shrimp groups (e.g., penaeids,
stenopodids) by the large second abdominal pleura that
overlap those of both the first and third somites; moreover, all carideans lack terminal chelae on the third
pereiopods (Fig. 1a). Astacideans (Fig. 2) include the
crayfishes of the northern and southern hemispheres
and the marine lobsters; only North American crayfishes are treated here. In contrast to the carideans,
crayfishes are rarely compressed laterally and their first
three pairs of pereiopods are always chelate. Extensive
body segmentation and the presence of jointed appendages on all metameres give most decapods a primitive appearance; yet the nervous, sensory, circulatory,
and digestive systems are complex, a necessary requirement for animals of large body size. The body of these
freshwater decapods is encased in an exoskeleton consisting of complex polysaccharides hardened with inorganic salts (except at joints, where the cuticle is thin
and pliable). The head and thoracic segments are fused
to form a large cephalothorax covered by a single
shield, the carapace (Figs. 1 and 2), but the six abdominal segments are individually distinct. The anterior portion of the cephalothorax contains a pair of large,
stalked eyes, a median rostrum, two pairs of antennae,
and a pair of mandibles. The body is composed of
somites, each with a pair of ventral, jointed appendages
that are serially homologous and basically biramous
(Fig. 3), but are modified for various functions (e.g.,
sensory, food handling, cleaning, gill-bailing, pinching,
walking, copulation, egg attachment and incubation,
and swimming). The typical biramous appendage is Yshaped and the base of the Y is attached to the somite.
The base is the protopodite (consisting of two joints —
coxopodite and basiopodite) that bears a mesial endopodite, typically of five podomeres, and a lateral exopodite, with few to many segments.
B. Organ System Function
The digestive system of crayfishes is simple and includes the mouth, a short tubular esophagus, the stomach, two large hepatopancreatic glands, a short midgut,
and a long tubular intestine extending dorsally through
the abdomen to the anus (Fig. 4). The relatively simple,
straight gastrointestinal tract has few significant areas
for storage or microbial degradation although the
stomach functions as a storage compartment (cardiac
portion), as a masticating structure (gastric mill), and
as a filter for gathering digestible material (pyloric portion). Morphological differences among species in the
grinding surfaces of the gastric mill relate to diet
(Caine, 1975). For example, the surface is reduced in
FIGURE 1 (A) Lateral view of generalized shrimp (after Hobbs and Jass, 1988). Ai, appendix interna;
Am, appendix masculina; As, antennal spine; B, basis; Bg, branchiostegal groove; Bs, branchiostegal
spine; Cp, carpus; D, dactyl; end, endopod; I. ischium; M, merus; P, propodus; and Sc, scaphocerite. (B)
Chela of second pereiopod with apical tufts of setae; Palaemonias ganteri (after Hobbs et al., 1977).
23. Decapoda
FIGURE 2
Dorsolateral view of generalized crayfish. Al, areola length; Ar, areola; As, antennal scale; Aw,
areola width; B, basis; Bcg, branchiocardiac groove; Bs, branchiostegal spine; C,carpus; Car, rostral carina; Cs,
cervical spine; D, dactyl of chela; I, ischium; Lru, lateralramus of uropod; M, merus; Mru, mesial ramus of
uropod; Ms, marginal spine; P, propodus-palm of chela; Pl, palm length; Ple, pleuron; Pocl, postorbital
carapace length; Pr, postorbital ridge; Ps, postorbital spine; Pw, width of palm; R, rostrum; Soa, suborbital
angle; Ter, tergum; and pleopods on ventral side of abdomen.
FIGURE 3
Biramous appendages:(a) Palaemonias alabamae, third maxilliiped (after Smalley, 1961); (b)
Cambarus (Depressicambarus) strigosus Hobbs, dorsal view of telson and biramous uropod (after Hobbs,
1981); and (c) Procambarus (Ortmannicus) lunzi (Hobbs), third pereiopod.
957
958
H. H. Hobbs III
FIGURE 4
Generalized diagram of crayfish, demonstrating internal organization.
some cave crayfishes that feed on organic silt and expanded in epigean species that utilize macromaterials.
The pH of the stomach is relatively alkaline and the enzymes of the gastrointestinal tract and hepatopancreas
are particularly effective in the processing of a wide
spectrum of food materials. Chymotrypsin and pepsin
are lacking, but an alkaline protease, trypsin, cellulase
(hepatopancrease origin), lysozyme (muramidase), and
probably chitinase and chitobiase are present (Brown,
1995).
The circulatory system is an open or lacunar system
and consists of a heart, arteries (no veins), and sinuses.
The heart is situated in a large middorsal pericardial sinus, and blood in the sinus enters the heart via three
pairs of ostia (valves). The nearly colorless blood is
pumped into arteries that distribute it to various organs
and afferent sinuses leading to the gills, and the blood
returns through a series of efferent sinuses that ultimately join to the pericardial sinus. Hemocyanin is the
principal blood pigment transporting oxygen.
The gills are attached to the thoracic appendages
(maxillipeds and pereiopods) and are situated along either side of the thorax in the bronchial (gill) chamber.
Lateral extensions of the carapace (branchiostegites)
cover the gills (Figs. la, 2, and 7). In crayfishes, anterior
to the gills, paddlelike second maxillae (scaphognathites, see Burggren and McMahon, 1983) beat back
and forth, drawing water over the gill filaments to ensure gas exchange. The gills of crayfishes (trichobranchs composed of unbranched, fingerlike filaments)
and shrimps (phyllobranchs consisting of platelike elements) are arranged in three series: the single
podobranchia borne on the coxa, the arthrobranchiae
on the articular membrane between the coxa and basis,
and the pleurobranchia borne on the pleura. The pleurobranch series is absent in all of the North American
crayfishes except in members of the genus Pacifastacus,
which have a single pair.
Considerable water constantly diffuses into the
blood through the gill surfaces of freshwater decapods.
Osmotic control and excretion are maintained by two
large antennal (green) glands situated ventral to the
subesophageal ganglia and anterior to the esophagus.
These glands have an intimate association with the circulatory system and thus maintain water balance
(Flynn and Holliday, 1994). The green gland consists
of a complexly folded end sac, a convoluted labyrinth,
a nephridial canal, a bladder, and an excretory pore situated at the coxae of the second antennae. Crayfishes
excrete a quantity of hypotonic urine (some ammonia,
urea, uric acid, and amines), which is generally isosmotic with the blood. High ammonia excretion rates
have been correlated with low pH values (pH 4.6; Daveikis and Alikhan, 1996).
The central nervous system functions as the destination for sensory input, the center of synaptic integration, and the source of motor control and consists of
paired supraesophageal and subesophageal ganglia,
which are joined, forming the “brain.” This juncture is
accomplished by a pair of connectives extending caudally from the subesophageal ganglia to the double
ventral nerve cord which has segmental ganglia connected by commissures (see Sandeman et al., 1992 for a
proposed common nomenclature for homologous
structures of the brain of crayfishes, crabs, and spiny
lobsters). Of significance, 30 – 40% of crayfish brain
volume is devoted to processing olfactory input (Mellon et al., 1992). Cuadras (1993) evaluated the eleven
categories of secretory organelles (membrane bound
carriers) present in the central nervous system of crayfishes. The peripheral nervous system consists of nerves
and receptor cells that function primarily as conducting
23. Decapoda
pathways (see Swerup and Rydqvist, 1992). Atwood
and Nguyen (1995) reviewed neural adaptations of
crayfishes to different activity levels, describing how
various species are equipped for a variety of environmental changes.
The basal segment of each first antenna supports
an organ of balance (statocyst). Chemoreceptors are
abundant on mouth parts, pereiopods, and antennae;
these last appendages also function as tactile organs.
Lateral rami of the antennules bear chemoreceptive
sensilla called aesthetascs that are innervated by sensory neurons. Tierney and Dunham (1984) and Tierney
et al. (1984) suggested that these structures may be receptive to pheromones although there have been conflicting results from various studies concerning under
what conditions and the location of receptor sites
where chemical sensation takes place (see Oh and Dunham, 1991; Dunham and Oh, 1992).
Rutherford et al. (1996) reported that in agonistic
encounters of conspecific Form I males (breeding
males — see below) of Orconectes (Procericambarus)
rusticus (Girard), winners had higher rates of antennal
movements than losers. Winners also demonstrated
longer periods of “large amplitude depression” (LAD —
holding antennules approximately 45° below the horizontal plane) which is used often in aggressive encounters. The researchers postulated that the increased use
of crayfish LAD movements may be in response to some
“stress pheromone” delivered to them by their opponent’s maxillipeds. Pheromones and other biochemicals
are used by crayfishes to recognize species, sex, food,
the presence of animals that are stressed, disturbed, or
physically damaged. These alarm substances are widespread and the detection of these will elicit distinct responses by crayfishes (Hazlett, 1994b).
Dunham et al. (1997) examined the use of the inner and outer rami of the antennules in mediation of
feeding behavior of Cambarus (Cambarus) bartonii
bartonii (Fabricius) and found that the several behavior
patterns used were dependent on the presence of either
inner or outer antennular rami, but not on both. Many
setae on the body surface contain sensory neurons and
are mechanoreceptors of one kind or another and at
least three sensory structures on the chelipeds appear to
have chemo- and mechanosensory function (Borash
and Moore, 1997). A pair of large compound eyes
characterizes most decapods [excluding stygobites (obligate, aquatic cave dwellers)], and reduced eye size in
most primary burrowing species); each eye is composed
of a number of virtually identical, discrete optical units
called ommatidia (“mosaic eyes” — good for detecting
movement, but limited resolving power) that are
arranged in geometrical array. In addition to compound eyes, a pair of neurons originating in the sixth
959
abdominal ganglion function as extraocular photoreceptors and are responsible for photonegative behavior,
but also receive information from mechanoreceptors in
the abdomen (see Fernández-de-Miguel and Aréchiga,
1992; Pel and Moss, 1996).
Further details of the morphology of decapods, in
general, can be examined in numerous invertebrate zoology textbooks (see also Holdich and Reeve, 1988),
and the external anatomy of crayfishes specifically is
treated well by Crocker and Barr (1968).
C. Aspects of Decapod Physiology
1. Respiration
The adaptation of organisms to various habitats
usually involves physiological, biochemical, and behavioral adjustments. Obviously a dynamic interaction exists between the external environment of an organism
and the functioning of its internal machinery. For example, reduced metabolic rates in cave decapods may
stem from biochemical and physiological adaptations
to greater environmental stability, to low number of
predators, or to a severely constrained resource base in
terms of quantity and variety (Huppop, 1985), as
demonstrated by whole animal respiration studies (see
Weingartner, 1977). Also, the reduction of oxygen consumption and energy turnover of gill tissues reported
by Dickson and Franz (1980) further substantiate the
highly specialized nature of physiological and biochemical adaptations in stygobitic animals.
In order for any life form to survive, the variety of
factors constituting the ambient environmental complex must not exceed genetically determined tolerance
limits for that species. That crayfishes and shrimps
have radiated into nearly every type of aquatic habitat
suggests that these crustaceans have been tolerant and
highly adaptable, particularly with regard to respiratory physiology. For example, Wiens and Armitage
(1961) demonstrated that, for the crayfishes Orconectes (Trisellescens) immunis (Hagen) and 0.
(Gremicambarus) nais (Faxon), the effect of body size
on oxygen consumption and the effects of both temperature and oxygen saturation on oxygen consumption
by them are highly significant (inverse relationship between body weight and metabolic rate; direct relationship between temperature and oxygen consumption).
They also showed that the cumulative effect of increased temperature and lowered oxygen concentration
produces greater stress than either parameter acting independently. 0. immunis has a daily metabolic rate
10% lower than 0. nais, and 0. nais does not regulate
as well as does 0. immunis when under stressed conditions. Therefore, the inability of 0. nais to maintain a
higher and more nearly regulated metabolic rate (as
960
H. H. Hobbs III
FIGURE 5
Cambarus tenebrosus from epigean and hypogean (stygophile) lotic habitats in the
midwestern and southeastern United States.
does 0. immunis) under such conditions, probably excludes it from roadside ditches where 0. immunis is
found. Hence, 0. immunis is better adapted physiologically to live under environmental conditions of periodic
high temperatures and low oxygen saturations (see
Reiber, 1997). Daveikis and Alikhan (1996) found that
Cambarus (Puncticambarus) robustus Girard from an
acidic, metal-contaminated lake in Ontario, Canada,
had lower oxygen consumption rates than conspecific
crayfishes from a circumneutral, uncontaminated, high
velocity stream. Nelson and Hooper (1982) showed
that the shrimp, Palaemonetes kadiakensis Rathbun,
exhibited an acute thermal preference within a temperature range of 6 – 10°C below their maximum tolerance
limits of 37 – 40°C. This shrimp was unable to tolerate
FIGURE 6
sustained exposure to temperatures greater than 32°C.
For additional treatment of metabolism, temperature
acclimation, preference, and avoidance in these two
groups of crustaceans, see McWhinnie and O’Connor
(1967), Crawshaw (1974), Becker et al. (1975), Loring
and Hill (1978), Mathur et al. (1982), Layne et al.
(1985), and Reiber (1995).
Burrowing crayfishes are often observed above the
water table; many species are capable of remaining out of
water for extended periods in the humid environment of
their tunnels (Hobbs, 1981; McMahon and Hankinson,
1993, and McMahon and Stuart, 1995). The same is true
for many cavernicolous crayfishes (stygophiles – Fig. 5,
and stygobites – Fig. 6), which, in the saturated air, can
move from one pool to another or remain out of the wa-
Orconectes pellucidus, a stygobitic crayfish from Kentucky and Tennessee.
23. Decapoda
ter for hours. Burrowers and surface-dwelling crayfishes
are commonly noted at the air – water interface, aerating
in response to low oxygen levels; the oxygen concentration in burrow waters is frequently below 2 mg / L
(Hobbs, 1981), and pool and ditches often become oxygen-depleted. These species that inhabit such oxygenpoor habitats have evolved a greater branchial volume by
adopting several characteristics (Figs. 7 and 8), such as a
vaulted and elongated carapace as well as branchiocardiac grooves that are closer together thus narrowing the
areola and raising the height of the chamber. Kushlan
and Kushlan (1980) observed the shrimp Palaemonetes
paludosus (Gibbes) swimming at the surface using oxygen that diffused across it as a means of surviving low
oxygen tension in the Everglades.
Many species (e.g., the crayfish Procambarus
(Girardiella) simulans (Faxon)) are oxygen regulators
and increase their ventilation rates in response to reduced oxygen or increased carbon dioxide concentrations. Others [e.g., Pacifastacus (Pacifastacus) l. leniusculus (Dana)] are oxygen conformers, lacking specific
adaptations for regulation of oxygen uptake at low oxygen tensions (see Vernberg, 1983). For further discussion
of respiratory physiology in crayfishes and shrimps, see
961
FIGURE 7
Semi-diagrammatic cross section through the thoracic
region of (a) Cambarus diogenes, a primary burrower living in
oxygen-poor groundwaters and (b) Procambarus(Pennides) versutus
(Hagen), dwelling in the oxygen-rich lotic environment (after Hobbs,
1976); arrows point to branchiocardiac grooves.
Moshiri et al. (1970), Cox and Beauchamp (1982),
Nelson and Hooper (1982), and Reiber (1995).
2. Ecdysis
Molting (ecdysis), a process critical for growth and
seasonal changes in form (dimorphic only in male cambarines), is a recurring crisis in the lives of shrimps and
FIGURE 8 Dorsolateral views of (a) Cambarus (Jugicambarus) gentryi Hobbs, a primary burrower (Cheatham
County, TN); (b) Procambarus lunzi, found in roadside ditch lentic environments (Hampton County, SC);
(c) Cambarus (Jugicambarus) distans Rhoades, a dweller of swift, surface streams (Dade County, GA); and
(d) Procambarus (Ortmannicus) enoplosternum Hobbs, occupies sluggish surface streams (Washington
County, GA).
962
H. H. Hobbs III
crayfishes. It involves far more than periods of discontinuous growth facilitated by a simple splitting of the
cuticle and secretion of a new exoskeleton. The life of a
decapod consists of alternating periods of premolt,
molt, postmolt, and intermolt; all of these phases are
controlled by hormones (endocrines). Neurosecretory
cells occur in the sensory papillae and in the X-organs
of the medulla terminalis; both secretary masses are situated in the eye stalk. The sinus gland, also located
within the eye stalk, receives secretions from the Xorgans and, in turn, produces hormones that inhibit
ecdysis. The Y-organs, a pair of glands in the maxillary
somites, secrete hormones derived from dietary cholesterol (three ecdysteroids: ecdysone; 25-deoxyecdysone;
and 3-dehydroecdysone) that stimulate molting (under
the control of the X-organ). During postmolt and intermolt, the X-organ / sinus gland complex liberates a neuropeptide molt-inhibiting hormone, thus regulating the
length of the intermolt period (see Lachaise et al.,
1993). Under appropriate external or internal conditions (light, temperature, loss of limbs — see Stoffel and
Hubschman, 1974), sinus gland hormones are not released and the Y-organ, no longer suppressed, secretes
the molt-initiating hormone. This acts on the epidermis
to initiate premolt activities that affect most body
parts. Glycogen reserves are increased and minerals
(e.g., calcium) are resorbed from the exoskeleton and
stored. With the secretion of an enzyme that softens the
cuticle at its base, it pulls away from the epidermal
cells (apolysis), stimulating the formation of a new epicuticle that is impervious to the molting enzyme. At the
termination of premolt, the old cuticle splits, the animal emerges from the old exoskeleton (exuvium) including shedding the cuticle covering the gills (see Andrews and Dillaman, 1993), and the soft body within
retains the maximum volume due to imbibing of water
by the decapod. Tissues grow rapidly; and during the
short postmolt period, the new layer becomes rigid as
the stored minerals are deposited in the hardening cuticle. Bendell-Young and Harvey (1991) found that crayfishes from acid lakes have carapaces with lower Mg
concentrations than those from circumneutral lakes,
suggesting that low pH conditions may prevent normal
mineralization during molting. After postmolt, the
crustacean may enter intermolt; however, complete intermolt probably occurs only in adults, and the rapidly
growing (molting) juveniles generally have, at best, a
very short period of stability (see Aiken and Waddy,
1992; Wheatly, 1995; Wheatly et al., 1996).
3. Reproductive Physiology
The development and function of gonads and the
ontogeny of secondary sexual characters in the dioecious decapods are regulated by hormones. The Y-organ
influences the development and maturation of gonads in
juveniles. In females, the sinus gland / X-organ complex
is important in controlling the ovary. During nonbreeding periods, the sinus gland produces a hormone that
inhibits egg development. However, during the breeding
season, the blood level of this hormone declines and egg
development is subsequently initiated. Development of
the testes and male sexual characteristics is controlled
by hormones produced in the androgenic gland, a small
mass of secretory tissue near the vas deferens (Taketomi
et al., 1996). Additional information concerning the reproductive cycle for shrimps and crayfishes is presented
below. Even though the source and nature of interspecific and intraspecific chemicals (pheromones) are unknown for crayfishes, studies have shown that these
chemical signals play an integral role in the behavior of
these decapods, although other studies have not demonstrated this (see review by Bechler, 1995). The role of
chemoreception in mediating agonistic behavior, species
recognition, and mate location in crayfishes has been
documented by Hazlett (1985). Discrimination between
male and female conspecifics using chemical cues has
been reported by some authors; these pheromones may
promote sex recognition and induce mating behavior
(see Hazlett, 1985; Dunham and Oh, 1996). Tierney et
al. (1984) determined that the site of pheromone reception in Orconectes (Crockerinus) propinquus (Girard) is
the lateral antennular flagellum (exopod) bearing the
chemoreceptive sensilla (aesthetascs).
4. Other Physiological Adaptations
As ectotherms, these decapods are subjected to
temperature changes that directly affect metabolic rate,
and most species demonstrate various thermal tolerances (most upper tolerances range from 32 to 36°C)
and preferences for their aquatic medium. Unfortunately, upper lethal temperatures are known only for a
few species (see Cox and Beauchamp, 1982, and references therein). Nelson and Hooper (1982) showed experimentally that Palaemonetes kadiakensis exhibited
an acute thermal preference (short-term response)
within a temperature range of 6 – 10°C below their
maximum tolerance limits. Taylor (1984) determined
for three southeastern United States crayfishes that
both the acute preference and final preferendum primarily represented the temperature that they encountered during daily or seasonal activity cycles. He hypothesized that temperature choice does not function in
temperature regulation except in the very broadest
sense; instead, it serves as an “error detection system,”
and cues for behavioral arousal from either seasonal hibernation or from their daily refugia. Clearly, water
temperature plays an important role in the ecology of
these crustaceans (see Mundahl and Benton, 1990).
23. Decapoda
Biological rhythms are recognized in decapods and
reviewed by DeCoursey (1983). Not only are organismic cycles pronounced (e.g., crayfishes have daily locomotor rhythms) but cellular and physiological rhythms
are prominently exhibited (e.g., circadian rhythmicity of
heart rate, Pollard and Larimer, 1977). Most seem to be
in response to environmental cues (e.g., light, temperature); however, even where rhythmic environmental input may be lacking, circadian clock regulation can be
maintained. For instance, Weingartner (1977) demonstrated a circadian activity pattern in the cave crayfish
Orconectes (Orconectes) i. inermis Cope from Indiana.
In a similar way, the Kentucky stygobite 0. (Orconectes)
pellucidus (Tellkampf) has retained a circadian clock
regulation of locomotor activity and oxygen consumption in spite of isolation from day – night cycles.
Crayfishes and shrimps are faced with the problems of constant influx of water by osmosis and loss of
salts by diffusion (Flynn and Holliday, 1994). Maintenance of constant body fluid composition is accomplished by active sodium uptake via the gills, production of a copious hypotonic (dilute) urine, and by some
reduction in boundary membrane permeability (see
Newsom and Davis, 1991; Dickson et al., 1991; Sarver
et al., 1994). As indicated above, the green gland is intimately associated with the circulatory system.
The chromatophore is an integument cell with
branched, radiating, noncontractile processes that are
associated with one or more types of pigment granules.
These white, red, yellow, blue, brown, or black pigments can flow into the processes or may be confined
to the center of the cell. A single chromatophore may
possess any number of pigments and is accordingly
classified as mono-, bi-, di-, or polychromatic (see
Fig. 9). In shrimps, the number of pigments and the
number of chromatophores can change (morphological
color change). However, physiological color change
FIGURE 9
Female Palaemonetes kadiakensis with eggs (note
chromatophores, e.g., small speckles on abdomen).
963
(rapid color adaptation to background resulting from
dispersal and concentration of pigments within the
chromatophores) is the more common type of color alteration and is controlled by hormones and the central
nervous system. Chromatophorotropins are released
from both the sinus gland and the central nervous
system. The latter elaborates several chromatophorotropins, resulting in pigments either dispersing or concentrating in the chromatophores (darkening or
blanching of body color).
Color morphs of shrimps and crayfishes have been
documented in a number of species, such as Macrobrachium rosenbergii (deMan), Orconectes propinquus,
Procambarus (Ortmannicus) a. acutus (Girard), Procambarus (Scapulicambarus) paeninsulanus (Faxon).
Studies have demonstrated that color variations may be
due to genetic differences (e.g., Black, 1975) or they can
be environmentally induced (see Fitzpatrick, 1987b;
Thacker et al., 1993).
Increasing awareness of human impact on the environment has directed attention to the effects of various
pollutants on the physiological ecology of crustaceans,
including shrimps and crayfishes. Accumulation of trace
metals occurs through runoff or direct dumping of
sewage or other effluents into aquatic systems. Lead has
become particularly important because of its relative
toxicity and increased environmental contamination via
automobile exhaust and highway runoff. Anderson
(1978) demonstrated for 0. (Gremicambarus) virilis
(Hagen) that exposure to lead resulted in damage to
gills, which caused a decrease in the uptake of oxygen;
ventilation rates increased as ambient lead concentrations increased. Hubschman (1967) noted that the toxic
effect of copper on crayfishes is dependent not only on
concentration and duration of exposure but also on the
age (or size) of the crustacean (see Maranhão et al.,
1995). At concentrations above 1 mg / L, respiratory enzymes are inhibited quite rapidly as the mechanism of
detoxification is apparently overwhelmed. At concentrations below 1 mg / L, copper has a chronic effect on
cell maintenance. For example, actively secreting cells
such as those of the antennal gland apparently are destroyed because cell repair cannot keep pace with cellular activity (see also Taylor et al., 1995). Cadmium, a
highly toxic metal, has been implicated as both a carcinogen and a mutagen and causes a number of cardiovascular problems (all resulting from enzyme dysfunction). Cadmium is bioaccumulated and thus when
incorporated in tissues of freshwater decapods can have
far-reaching effects within aquatic ecosystems as well as
in humans (e.g., Dickson et al., 1982; Thorp and Gloss,
1986; Bendell-Young and Harvey, 1991; Naqvi and
Howell, 1993a, b). Vermeer (1972) noted high mercury
levels in muscle tissue of 0. virilis from several localities
964
H. H. Hobbs III
in Manitoba and Ontario and suggested that this
species is a good indicator of mercury contamination in
aquatic ecosystems (see also Wright and Welbourn,
1993). Insecticides (Albaugh, 1972), herbicides (Naqvi
and Leung, 1983), lampricides, and numerous other
chemicals have been shown to have various detrimental
effects on freshwater decapods.
The bioaccumulation of metals and other toxic or
persistent chemicals poses particular problems for longlived cavernicoles (stygobites)(see below, Cooper and
Cooper, 1978). Dickson et al. (1979) showed higher
concentrations of cadmium and lead in tissues of Orconectes (Orconectes) a. australis (Rhoades) (a stygobite) than in those of Cambarus (Erebicambarus) tenebrosus [a stygophile — this species now inclusive of C. cahni
(see Hobbs, 1989), C. laevis, and C. ornatus (see Taylor,
1997)], crayfishes occurring syntopically in a Tennessee
cave. It is possible that some stygobitic decapod populations could be extirpated not from a single, massive dose
of a chemical (though this has occurred; see Bechler,
1983; Hobbs III, 1987), but slowly from long-term exposure to low levels of these metals or other toxic substances. Because the longevity of primary burrowing
species is apparently greater than that for nonburrowing
taxa living in surface waters, a similar threat from
chronic pollution exists for these crustaceans. This is
particularly true for crayfishes living in roadside ditches
that are periodically sprayed with various pesticides
(Hubschman, 1967; Dimond et al., 1968).
III. ECOLOGY AND EVOLUTION
A. Diversity, Distribution, and Evolutionary
Relationships
Most freshwater decapods of North America occur
in one or more of the following habitats: shallow
(1 – 2 m depths, although this is probably a function of
sampling bias) lentic and lotic waters, epigean and hypogean habitats, oligotrophic to hypereutrophic lakes,
ponds, marshes, ditches, steep-gradient headwater
streams, low-gradient large rivers, springs, stygean
streams and pools, and terrestrial burrows leading to
groundwater. Individuals have been found in lakes or
springs at depths of 30 – 90 m (stygobitic crayfishes in
submerged caves in north-central Florida are observed
by divers at depths greater than 30 m) but crayfishes
certainly thrive in shallow, littoral areas. They are most
abundant in the central, eastern, and southern regions
of the United States, while a few species are found in
the Pacific slope drainages; they are rare or absent in
the large area of the western Great Plains and the
Rocky Mountains (see Taylor et al., 1996). This simply
means that American crayfishes are crustaceans possessing biological plasticity. They have become established in nearly every type of freshwater habitat, except
glacial and thermal effluents, and some can tolerate
brackish water. Considerable data suggest that habitat
has a channelizing influence (convergence) on certain
morphological aspects of crayfishes (Hobbs, 1976). For
example, stygobites (a polyphyletic group) are characterized by an albino-sightless combination and attenuated appendages; they are classic K-strategists and
demonstrate extreme resource efficiency. In contrast,
“burrowers” (also polyphyletic) possess short, broad,
flattened chelipeds, lack carapace spines, have a reduced abdomen and tail fan (length, width, and
height), and possess an enlarged gill chamber (an adaptation to the oxygen-poor burrow habitat — see above)
(Fig 7). Water currents have certainly influenced the
body form of crayfishes, with those species dwelling in
fastest flowing water having depressed (dorsoventrally
flattened — most common) or compressed (laterally deflated) bodies; these include, for example, Orconectes
(Gremicambarus) compressus (Faxon) and Procambarus (Tenuicambarus) tenuis Hobbs. Spination is also
typically reduced in such species. Those crayfishes inhabiting riffle areas of streams and residing in pebble
and cobble substrates (rubble dwellers) are markedly
depressed dorsoventrally; examples of such taxa are
Cambarus (Jugicambarus) friaufi Hobbs and Cambarus
( J.) brachydactylus Hobbs.
Several decapods (Infraorder Brachyura — crabs)
occur sporadically (naturally or introduced) in coastal
brackish or freshwater systems, but are not included in
the following discussions, numerical summaries, or key.
They are represented by the portunid blue crab, Callinectes sapidus Rathbun, a xanthid mud crab,
Rhithropanopeus harrisii (Gould) (found in streams of
the Atlantic and Gulf coasts and introduced along the
coasts of California and Oregon), and a grapsid river
crab, Platychirograpsus spectabilis Rathbun, introduced from eastern Mexico into the Hillsboro River
near Tampa, Florida (Powers, 1977). Of curious interest is the occurrence of the grapsid crab, Hemigrapsis
estellinensis Creel, in Estelline Salt Spring in Hall
County, Texas. This representative of the otherwise
marine family Grapsidae is endemic (probably a Pleistocene relict) to the spring which is located 800 km
from the sea at an elevation of 531 m (see Reddell,
1994). For further information on cavernicolous crabs,
refer to Guinot (1994).
1. Shrimps
The atyid shrimps (fingers of chelate pereiopods
with apical tufts of setae, Fig. 1B) of North America
are restricted to two genera, one with two southeast-
23. Decapoda
ern, obligate cave species and the other represented by
two species found in coastal streams of California.
Freshwater palaemonids (fingers of chelate pereiopods
lacking apical tufts of setae, Fig. lA) are a very successful family of shrimps; the genus Palaemonetes contains
approximately 17 species in the western hemisphere,
and Macrobrachium is assigned about 100 species
worldwide (35 in the western hemisphere). These genera are represented in North America by 10 and 12
species, respectively. Shrimps are generally found in the
macrophyte-rich littoral zone of lakes or in sluggish,
vegetation-choked streams. Within these two families
residing in the confines of the United States, five
shrimps are stygobitic and are very restricted in distribution (two species known to occur only in a single
karst locality).
The North American stygobitic decapods (inclusive
of Bahamas, Barbuda, Belize, Bermuda, Caicos Islands,
Canada, Cuba, Dominican Republic, Guatamala, Hispaniola, Honduras, Isla de Pinos, Isla Mona, Jamaica,
Mexico, Puerto Rico, and the United States) are represented by 36 shrimps assigned to five families and 16
genera:Agostocarididae [Agostocaris (2)], Alpheidae
[Automate (1), Potamalpheops (1), Yagerocaris (1)],
Atyidae [Palaemonias (2), Typhlatya (8)], Hippolytidae
[Barbouria (1), Calliasmata (1), Janicea (1), Somersiella
(1)], and Palaemonidae [Cryphiops (2), Creaseria (1),
Macrobrachium (3), Neopalaemonon (1), Palaemonetes (3), Troglocubanus (6)], Procarididae [Procaris (1)]
(Hobbs III, 1998).
a. Atyidae North American freshwaters are inhabited by three federally endangered and one probable extinct atyid species:Palaemonias ganteri Hay, P. alabamae Smalley, Snycaris pacifica (Holmes), and S.
pasadenae (Kingsley), respectively (Hedgpeth, 1968).
The first two taxa are stygobitic shrimps, the former restricted to Mammoth Cave in Kentucky, and the latter
is known only from five caves (three groundwater
basins) in Madison County, northeastern Alabama (Jacobson and Hartfield, 1997; McGregor et al., 1997).
Because most atyids currently inhabit warmer southern
aquatic ecosystems, it is probable that these two stygobites are thermophilic relicts of a former, more widely
distributed common ancestor. Syncaris pacifica and S.
pasadenae are found in small coastal streams of California. The former is reported from Marin, Napa, and
Sonoma counties and the latter is previously known
from Los Angeles, San Bernardino, and San Diego
(may be erroneous report) counties but is likely extinct
due to habitat destruction and over-collecting.
b. Palaemonidae Those North American palaemonids that occur within the United States and that
965
FIGURE 10 Geographical distribution of Palaemonetes kadiakensis
and P. paludosus.
are strictly freshwater inhabitants are represented by
10 species belonging to the genus Palaemonetes (three
are stygobitic): Palaemonetes antrorum (Benedict)
from wells and caves in Hays and Uvalde counties,
Texas (federally endangered), P. holthuisi (Strenth)
from a cave in Hays County, Texas (the only known
site in the United States where two species of stygobitic
shrimps are found together), and P. cummingi Chace
from a cave in Alachua County, Florida. Palaemonetes
cummingi is listed as “vulnerable” (threatened) on the
1996 IUCN Red List of Threatened Animals (Baillie
and Groombridge, 1996). Seven are inhabitants of
lentic and sluggish lotic systems, including springs (see
Strenth, 1976, 1994; Wu and Brown, 1980) (Fig. 10 —
distribution map does not include disjunct populations
such as P. paludosus in Tygart Valley River, Randolph
County, West Virginia — population probably introduced). In addition, five species assigned to the widespread genus Macrobrachium (North, Central, and
South America, and the West Indies) are found in
freshwaters in the middle and lower Mississippi River
drainage and in low-lying streams from New Jersey to
Texas (Figs. 11 and 12). Macrobrachium acanthurus
(Wiegmann) ranges from Georgia to Brazil (also Bahamas) and is found in the United States from Georgia
and Florida to Texas along the Gulf coast. Macrobrachium carcinus (Linnaeus) is the largest and most
spectacular species and is found from Florida to Texas
and south to Brazil and the West Indies. Macrobrachium ohione (Smith) is the only endemic species of
the genus in North America and is found from Virginia to Texas and in the lower and middle Mississippi
River drainage (Fig. 11). The natural range of M.
olfersii (Wiegmann) is from Mexico to Brazil (Chace
966
H. H. Hobbs III
FIGURE 11 Geographical distribution of Macrobrachium ohione.
Gross reduction in populations throughout its range is attributable to
loss of suitable habitat.
and Hobbs, 1969); however, it likely has been introduced into the St. Augustine, Florida area (Hedgpeth,
1949). As referred to above, the giant Malaysian
prawn, M. rosenbergii, has been introduced into ponds
of central Florida, South Carolina, and numerous
FIGURE 12
other parts of the world (e.g., Israel, Jamaica) (Smith
et al., 1978).
Strenth (1976), in a hypothetical discussion of the
origin and dispersal of Palaemonetes and Macrobrachium, proposed that freshwater Palaemonetes may
have arisen during the late Mesozoic or early Cenozoic
and that the widely disjunct species of the genus are
largely of monophyletic or of limited polyphyletic
origin. He presented salinity tolerance data from laboratory experiments supporting the premise that worldwide dispersal of freshwater Palaemonetes over large
expanses of open ocean would have been unlikely. Results of his studies on P. kadiakensis, coupled with
those of others for P. paludosus, demonstrate that this
group of shrimps cannot osmoregulate in salinities
greater than 25 ppt. Thus, the distributional patterns of
Palaemonetes and Macrobrachium appear consistent
with the theories of plate tectonics and continental
drift. He proposed that Palaemonetes preceded Marcobrachium into the Gulf of Mexico area and that Macrobrachium (primarily juveniles), through competitive
exclusion, barred Palaemonetes from the middle latitudes. Most of the freshwater species of Palaemonetes
are found above or below the 30° latitudes; or, if found
between, they always occur at higher elevations or are
Geographical distribution of Macrobrachium acanthurus (filled circles), M .carcinus (filled
squares), and M. olfersi (filled triangles) (after Hedgpeth, 1949).
23. Decapoda
located at considerable distances from the coast. Parameters that appear to limit the further spread of Macrobrachium are current velocity, salinity, temperature
tolerances, and anthropogenic perturbations.
2. Crayfishes
The freshwater astacideans (superfamily Astacoidea) constitute a very successful group and currently
are represented by 393 described species and subspecies
(4 extinct species), 342 species and subspecies which
are restricted (introductions ignored) to North America, north of Mexico (see Hobbs, 1986 for lineages of
the major groups in North America). These are assigned to twelve genera and two families (Table I), of
which species richness sustains a progressive decline
with increasing latitude from 30 – 60°N (France, 1992).
The families, Astacidae and Cambaridae, have historically occupied allopatric ranges. The former inhabits
the Pacific slope drainages, except for Pacifastacus
(Hobbsastacus) gambelii (Girard) which has crossed
the divide into the upper Missouri; and the Cambari-
TABLE I
967
dae occurs in some Hudson Bay watersheds, the Atlantic, and Gulf drainages from southern Canada to
Honduras, certain Pacific slope basins in Mexico, and
Cuba (Fig. 13). The three most successful genera of
crayfishes are Procambarus, Cambarus, and Orconectes (166, 86, 81 species and subspecies, respectively — see below). The coastal plain of the southern
states is dominated by Procambarus, whereas members
of the genus Cambarus (Fig. 13) are major components
of aquatic environments from the Cumberland Plateau
to the North and East. West and northwest of the
Cumberland Plateau, species of the genus Orconectes
are the most commonly encountered crayfishes. Hobbs
(1989) produced a useful listing of the crayfish faunas
by country and state and included references treating
species within each state. Two recent publications that
discuss decapods in the northeastern United States are
those of Smith (1988) and Peckarsky et al. (1990), and
the following contemporary publications present data
for decapods within various states and Canada:Guiasu
et al. (1996) — Ontario, Canada; Deyrup and Franz
List of Freshwater Decapod Genera and Subgenera Occurring in the United States
Infraorder
Family
Subfamily
Genus
Subgenus
N. A.
U. S.
Caridea (Shrimps)
Atyidae
————
Palaemonidae
————
————
Palaemonias*
Syncaris
Palaemonetes*
Macrobrachium
————
————
————
————
Astacidae
————
————
Cambarellinae
Pacifastacus
Pacifastacus
Cambarellus
Cambarellus
Cambarellus
Barbicambarus
Bouchardina
Cambarus*
Hobbseus
Pacifastacus
Cambarellus
Dirigicambarus
Pandicambarus
————
————
Aviticambarus*
Cambarus
Depressicambarus
Erebicambarus*
Exilicambarus
Glareocola**
Hiaticambarus
Jugicambarus*
Lacunicambarus
Puncticambarus*
Tubericambarus
Veticambarus
Distocambarus
Fitzcambarus
Creaserinus
Fallicambarus
————
————
4
2
10
12
28
5
3
9
1
7
1
1
3
7
16
5
1
1
9
23
4
15
1
1
2
3
9
7
4
7
4
2
6
5
17
5
3
0
1
7
1
1
3
7
16
5
1
1
9
23
4
15
1
1
2
3
9
7
4
7
Shrimp Totals
Astacidea (Crayfishes)
Cambaridae
Cambarinae
Distocambarus
Fallicambarus
Faxonella
Hobbseus
(Continues)
968
TABLE I
Infraorder
H. H. Hobbs III
(Continued)
Family
Subfamily
Genus
Subgenus
N. A.
U. S.
Orconectes*
Billecambarus
Buannulifictus
Crockerinus
Faxonius
Gremicambarus
Hespericambarus
Orconectes*
Procericambarus
Rhoadesius
Tragulicambarus
Trisellescens**
Acucauda
Austrocambarus*
Capillicambarus
Girardiella
Hagenides
Leconticambarus*
Lonnbergius*
Mexicambarus
Ortmannicus*
Paracambarus
Pennides
Procambarus
Remoticambarus*
Scapulicambarus
Tenuicambarus
Villalobosus
————
1
6
14
3
5
7
8
25
2
1
9
1
20
3
19
10
14
2
1
56
2
19
1
1
6
1
10
1
393
1
6
14
3
5
7
8
25
2
1
9
1
1
3
18
10
14
2
0
50
0
18
0
1
5
1
0
1
342
Procambarus*
Troglocambarus*
Crayfish Totals
Summary listing: [refer also to Hedgpeth (1949), Strenth (1976), and Hobbs (1989)].
Infraorder Caridea (shrimps)
Family Atyidae
Genus Palaemonias — 4;
Genus Syncaris — 2;
Family Palaemonidae
Genus Palaemonetes — 10;
Genus Macrobrachium — 12;
Infraorder Astacidea (crayfishes)
Family Astacidae
Genus Pacifastacus — 2 subgenera and 8 species;
Family Cambaridae
Subfamily Cambarellinae
Genus Cambarellus — 3 subgenera, 17 species;
Subfamily Cambarinae
Genus Barbicambarus — monotypic;
Genus Bouchardina — monotypic;
Genus Cambarus — 12 subgenera and 86 species;
Genus Distocambarus — 2 subgenera and 5 species;
Genus Fallicambarus — 2 subgenera and 16 species;
Genus Faxonella — 4 species;
Genus Hobbseus — 7 species;
Genus Orconectes — 11 subgenera and 81 species and subspecies;
Genus Procambarus — 16 subgenera and 166 species and subspecies; only 12 subgenera and 124 species and subspecies in North
America north of Mexico [Procambarus (Austrocambarus) primaevus (Packard) is an extinct species known only from tertiary
deposits in Wyoming and is not included in these figures];
Genus Troglocambarus — monotypic.
Respective numbers of species in North America (inclusive of Belize, Canada, Cuba, Guatemala, Honduras, Isla de Pinos, Mexico, and the
United States) are included. Data are presented only for those genera that are assigned U. S. species (* denotes genera/subgenera having stygobitic species; ** see Bouchard and Bouchard, 1995).
23. Decapoda
969
FIGURE 13
Geographical distribution of North American crayfish genera (not inclusive of
introductions) (modified from Hobbs, 1988).
(1994) and Franz et al. (1994) — Florida, Georgia; Page
(1985) — Illinois; Page and Mottesi (1995) — Indiana;
Martin (1997) — Maine; Pflieger (1996) — Missouri;
Cooper and Braswell (1995) — North Carolina; Eversole (1995) — South Carolina; Jezerinac et al. (1995) —
West Virginia; and Hobbs III and Jass (1988) — Wisconsin. The reader should refer to Taylor et al. (1995)
for a more complete listing of publications having state
crayfish data.
Many epigean crayfishes occur abundantly in large
interstices that afford protection and concealment (dark
thigmotactic cover; Alberstadt et al., 1995), particularly
during the daytime hours, such as among stones, weed
beds, leaf litter, brush piles, logjams, roots of riparian
trees; and they often find shelter in cans, tires, or among
other anthropogenic debris. The genus Cambarellus in
North America is distributed below the Fall Line zone
in the Gulf Coastal Plain. Whereas a few species are
known only from lotic habitats, most members of this
genus, as well as those of Faxonella and Bouchardina,
frequent lentic habitats such as ponds, backwaters of
streams, and roadside ditches. These small crayfishes
are successful in littoral waters subject to low oxygen
concentrations and elevated temperatures. The
970
H. H. Hobbs III
FIGURE 14 Pool habitat for the stygobitic crayfish Orconectes a. australis in Russell Cave, Jackson County, AL.
members of the genus Hobbseus also occupy such habitats, but they are more common in very small, shallow
streams. The monotypic Bouchardina probably lives in
similar habitats since it has been collected from leaf
litter and aquatic vegetation in a single stream backwater ditch. The monotypic Barbicambarus is found only
under large limestone rocks in the swift currents of
large streams in the Green and Barren river basins of
the Highland Rim. Of note, introductions of nonnative
species into various parts of the world are leading to
species displacements among crayfishes (see below;
Hobbs III et al., 1989; Olson et al., 1991; Clancy,
1997).
Hypogean species are represented by burrowing
and cave-dwelling crayfishes. Primary burrowers,
which are found in the genera Cambarus, Distocambarus, Fallicambarus, and Procambarus (Fig. 13), construct tunnels (Fig. 22a, d) in such habitats as seepage
areas, along stream banks, in open fields, and in other
low-lying areas. Among the stygobitic species are members of the genera Cambarus, Orconectes, Procambarus, and Troglocambarus; some of these cavernicolous crayfishes occupy lotic habitats while others
appear to have become adapted to lentic situations.
Those species inhabiting caves with active streams generally are found in pooled areas where they have aban-
doned their preferred, and now less abundant, cover of
cobble-gravel in favor of accumulated allochthonous,
food-rich, silty substrates (Hobbs III, 1976) (Fig. 14).
Some stygobitic crayfishes living in sluggish lotic / lentic
systems (e.g., caves developed in the porous Ocala
limestone of northcentral Florida) are found near cave
entrances where organic debris is available; they are
generally positioned on the walls and silty substrates of
these submerged solution passages (Hobbs et al., 1977;
Hobbs and Franz, 1986). Franz and Lee (1982)
showed that the geographical and ecological distributions of the cavernicolous crayfishes of Florida are correlated with low or high energy (food) cave systems.
One species, Troglocambarus maclanei Hobbs, is often
found on walls or clinging pendant to ceilings of submerged caves. The stygobitic crayfishes are derived
from surface ancestors that entered caves during late
Tertiary (Hobbs et al., 1977; see also Hobbs and Franz,
1986; Holsinger, 1988; Hobbs III, 1994, 1999).
a. Astacidae A single genus, Pacifastacus, is represented by eight species and subspecies in the Pacific
drainages of western North America and headwaters of
the Missouri River in Wyoming (Fig. 13). One species,
P. (Hobbsastacus) chenoderma (Cope), is extinct,
known only from Miocene and Pliocene (?) deposits,
23. Decapoda
and is not included in Table I. P. (H.) nigrescens (Stimpson) is probably extinct and P. (H.) fortis (Faxon) populations have declined significantly (Light et al., 1995)
and it is listed as a Federal Endangered Species.
These crayfishes are found in lentic and lotic habitats ranging from alpine lakes and streams to streams
crossing cold deserts. They occur in California, Idaho,
Montana, Nevada, Oregon, Utah, Washington,
Wyoming, and British Columbia (Fig. 13). Hobbs
(1974) suggested that two evolutionary lines arose
from a Mesozoic marine nephropoid ancestral stock.
One line (represented today by the genera Astacus,
Austropotamobius, and Pacifastacus) moved into the
freshwater habitat and maintained the primitive characteristic of absence of cyclic dimorphism in males, and
females exhibited a sclerite (annular plate) lacking both
a sinus and fossa. A second, less conservative stock also
made the transition to freshwaters (represented by extant members of the family Cambaridae). These males
evolved cyclic dimorphism and associated characters
(e.g., hooks on ischia of certain pereiopods in males),
while the females developed an annular sclerite with a
sinus or fossa for sperm storage (the annulus ventralis).
b. Cambaridae As previously stated, the ancestral
stock, in which cyclic dimorphism became established,
moved into freshwater habitats and gave rise to the
Cambaridae. This family consists of three subfamilies,
two of which (Cambarellinae and Cambarinae) occur
in North America (Fig. 13). The third, Cambaroidinae,
is restricted to eastern Asia-Amur Basin, Korea, and
Japan, and is treated only briefly in further discussions.
The following scenario is an abbreviation of arguments presented by Hobbs (1969, 1981, 1989) concerning the origin and dispersal of cambarid decapods.
Whether or not the Asian Cambaroidinae and the
American subfamilies shared a common freshwater ancestor or whether there were separate Asiatic and American invasions of freshwaters may never be known.
Hobbs, however, suggested that the ancestral cambarine
stock entered freshwaters of the southeastern portion of
the North American continent during the late Cretaceous or early Cenozoic eras. Through adaptive radiation, this ancestral Procambarus stock moved through
estuarine environments and populated available habitats in many of the streams flowing from the southern
and southeastern slopes of the existing continent. Except for rare passive dispersal resulting from stream
piracy or from crayfishes moving during periods of low
salinity from one stream mouth to another, stocks became isolated in their respective drainage systems.
Before the close of the Miocene (at least 25 million
years ago), some stream-dwellers were able to exist in
backwaters of lotic habitats; these forms gave rise to the
971
ancestors of some of the lentic species of Procambarus
(e.g., subgenera Ortmannicus and Scapulicambarus).
These crayfishes moved into an ecological vacuum and
were no longer confined to flowing water systems. Pools
may have dried, or oxygen concentrations became low,
or perhaps individuals simply moved across land from
one aquatic habitat to another. Whatever the reason,
additional ponds, lakes, ditches, streams, etc., were occupied and thus dispersal into lower elevation habitats
occurred; also several stocks penetrated the water table
by burrowing. Distocambarus occupied the groundwater niche and probably represents a remnant of a much
more widely distributed Piedmont stock. The monotypic Troglocambarus represents an offshoot of the Procambarus stock that moved into the karst groundwaters
in northcentral Florida [see below, and Hobbs et al.
(1977) for further discussion of the evolution of stygobitic decapods]. The genus Cambarellus probably arose
from a primitive ancestral Procambarus stock in the
Gulf coastal plain and occupied ephemeral swamps,
ponds, and ditches.
One of the lotic ancestral procambarid stocks
moved upstream, reaching the vicinity of the Cumberland Plateau, where ancestors to two very successful
genera, Cambarus and Orconectes, became established.
The major evolutionary lines were probably established
by the Miocene Epoch, and the final stage was set for
the refinement of evolutionary patterns begun during
the early Cenozoic. Such major features as large rivers,
the Appalachian and Ozark mountains, the coastal
plain, the Cumberland Plateau, the Nashville Basin,
and the Highland Rim influenced the dispersal of these
stocks. Superimposed on these were the effects of Pleistocene glaciation.
Representatives of the ancestral Cambarus stock
were successful in various habitats and spread through
lowland areas, giving rise to the species currently assigned to Fallicambarus. This genus, which probably
arose on the west Gulf coastal plain, developed into
primary burrowers. Barbicambarus is another descendant of the early Cambarus; it moved into sizeable
streams with large substrates, rapid currents, and high
oxygen concentrations. Hobbseus also may have
evolved from the cambaroid line (Fitzpatrick, 1977),
but it is more likely that it arose from the orconectoid
line [Hobbs (1969)].
The genus Orconectes probably evolved in the Interior Lowland Plateau province (Kentucky / Tennessee)
and radiated virtually in every direction (see Fitzpatrick,
1986, 1987a; Fetzner, 1996). Some orconectoid stock
successfully moved into the lowland regions where
species arose that are now assigned to the genus Faxonella. This genus also seems to have evolved on the
west Gulf coastal plain and today these small, tertiary
972
H. H. Hobbs III
burrowers are found in lentic situations (ponds,
ditches). They occur in flood plains and, when the water level rises, are fairly common in the shallow flowing
water. Bouchardina also is derived from the orconectoid
stock and occupies backwater areas. Ancestors of the
members of these two genera likely arrived along the
margins of the retreating Mississippi embayment during
the early Cenozoic.
Thirty-eight stygobite crayfish species and subspecies (refer to Table I) belonging to the family Cambaridae are assigned to the four genera:Cambarus (11
species), Orconectes (7 species and subspecies), Procambarus (19 species and subspecies), and the monotypic Troglocambarus. These obligate cave shrimps and
crayfishes were derived from surface ancestors that
made numerous colonizations of karst waters during
late Tertiary. Fluvial barriers would generally have had
minimal effects on the dispersal of stygobites (see Barr
and Holsinger, 1985, and Holsinger, 1988; also see
above and Hobbs et al., 1977 and Hobbs III, 1994 for
a more detailed discussion of their evolution and dispersal).
The profound effects of Pleistocene glaciation on
the distribution of organisms should be mentioned. The
obliteration of immense drainage systems (e.g., Teays)
resulted in total habitat destruction for virtually all organisms. The principal zoogeographical effects were
the elimination of populations and entire species from
the northern portions of North America, with displacements of species farther south than they had occurred
preglacially. Retreat of the glaciers resulted in the establishment of new drainage systems (e.g., the Ohio)
which undoubtedly were rapidly invaded (re-invaded)
primarily from the south and southeast, though some
species may have entered from more than one
refugium.
(1). Cambarellinae The subfamily Cambarellinae is
confined to North and middle America where its members are found in the coastal plain from northern
Florida to southern Illinois and Texas (Fig. 13) and,
disjunctly, on the Pacific slope and Central Plateau of
Mexico. Seventeen species are assigned to three subgenera in the single genus Cambarellus (Fitzpatrick, 1983);
however, only eight of them are found north of the Rio
Grande (Table I). In the United States, eight species are
known from the following states:Alabama, Arkansas,
Florida, Illinois, Kentucky, Louisiana, Mississippi, Missouri, Tennessee, and Texas (Fig 13); they may have
been introduced by humans into Georgia.
(2). Cambarinae The subfamily Cambarinae,
which is comprised of 10 genera and 368 species,
ranges from New Brunswick across southern Canada
to Mexico, Guatemala, Honduras, and Cuba (Fig. 13).
Ten genera represented by 326 species occur in North
America, north of Mexico (Table I; see Hobbs, 1989
for detailed presentation of genera, subgenera, species,
and subspecies of American crayfishes).
B. Reproduction and Life History
1. Shrimps
Like all astacideans, freshwater shrimps have external fertilization, with the ova being fertilized as they
are extruded and then attached to the pleopods of females. The two common epigean, freshwater shrimps,
Palaemonetes kadiakensis (Fig. 9) and P. paludosus,
have similar life histories, both species living approximately one year (Fig. 15). Although many individuals
reproduce only once (semelparous), Nielsen and
Reynolds (1977) observed ovigerous females of P. kadiakensis from Missouri also bearing mature eggs, indicating at least two broods of young per female
(iteroparous); Beck and Cowell (1976) also noted that
P. paludosus spawned twice. The length of the breeding
season varies with latitude and is generally longer in
southern localities. For instance, ovigerous females occur in populations of P. kadiakensis in Illinois and
Michigan from April until August, but are present in
Louisiana populations from February to October;
ovigerous females of P. paludosus occur throughout the
year in the Florida Everglades. As a general rule, breeding does not begin until late spring or early summer in
more northern areas. Only large females breed early in
the season, and they are replaced by smaller females as
the reproductive season progresses. Females are larger
and usually more abundant than males. Mature (eggbearing) females range from 20 – 49 mm in total length
and produce one or two broods (two is particularly
representative of populations of P. paludosus in Florida
where the growing season is longer). The number of
eggs produced per female is highly variable, ranging
from 8 to 160, with fecundity generally a linear function of total length. The incubation period depends on
water temperature and varies from 12 to 24 days;
zoeae larval lengths are approximately 4 mm at hatching and these free-swimming larvae pass through six
stages (varies from 3 to 8) in about three weeks. Accounts of larval development and postembryonic
growth in these two species are summarized by Hubschman and Rose (1969), Beck and Cowell (1976),
Beck (1980), Kushlan and Kushlan (1980), Page
(1985), and Hobbs III and Jass (1988). Shrimp usually
mature when they attain a total length of 20 mm although this is not necessarily the case. Most shrimp die
after reproducing, but some females molt within three
days (generally within 24 h) after young are released
and can later produce another brood. Generally, adults
disappear from the population in late summer or early
23. Decapoda
FIGURE 15
973
Generalized life history of Palaemonetes (see text for explanation).
fall, having a life span of approximately one year. Marine and brackish water Palaemonetes (e.g., P. intermedius Holthuis, P. pugio Holthuis) usually have
smaller and more numerous eggs per female than do
freshwater species. The production of fewer but larger
eggs may enable the freshwater larvae to hatch at an
advanced stage with a corresponding reduction in the
number of free-swimming larval stages.
Dobkin (1971) described three larval stages in the
stygobite, P. cummingi, and studies of atyids consist of
observations made on Palaemonias ganteri in Mammoth Cave, Kentucky (Hobbs et al., 1977; Leitheuser et
al., 1985); the Alabama cave shrimp, P. alabamae, conducted by Cooper (1975) and McGregor et al. (1997);
and Syncaris pacifica (Eng, 1981). The study of the life
history of Macrobrachium ohione has not received as
much attention as have the two taxa of Palaemonetes
previously discussed. This species is larger (total length
of ovigerous females ranging from 27 to 93 mm) and it
lives a maximum of two years (Fig. 16). [Huner (1977)
determined that M. ohione at Port Allen, Louisiana
lived up to two years.] Fecundity is relatively enormous,
the number of eggs varying from 6273 – 24,000 per
female (Truesdale and Mermilliod, 1979). Ovigerous females are known only during May in Illinois, from
March through September in Louisiana, and during
March in Texas. Generally, females are larger than
males and only the largest individuals in the population
bear eggs. Additional ecological data can be obtained
from Truesdale and Mermilliod (1979), Page (1985),
and references therein.
2. Crayfishes
Surprisingly few crayfishes have been studied carefully for extended periods. Although considerable data
are available for many species (e.g., Penn, 1943;
Fielder, 1972; Hamr and Berrill, 1985; Corey, 1990;
Mitchell and Smock, 1991; Norrocky, 1991; Johnston
and Figiel, 1997; see also Hart and Clark, 1987), most
are observations from isolated collections. Astacid lifehistory investigations are limited to Pacifastacus leniusculus leniusculus and Pacifastacus (Pacifastacus) leniusculus trowbridgii (Stimpson) (see references in Mason,
1970a, b; McGriff, 1983a). Life-history studies of the
more diverse cambarids have been conducted on only
about 20 species.
In a general sense, the life history of a crayfish can
be divided into several phases (Momot, 1984), each
having a slightly different component important to the
completion of the life history of a given cohort as well
974
H. H. Hobbs III
FIGURE 16
Generalized life history of Macrobrachium ohione (see text for explanation).
as to the long-term success of the species. Recently
hatched juveniles search lake littoral and stream riffle
areas for the best food and shelter available (see Figiel
et al., 1991). Rapid growth of young-of-the-year
(YOY) allows a partial escape from the devastating effects of predation (Gowing and Momot, 1979). As juveniles grow larger, they abandon littoral and riffle areas for deeper waters, only to return after reaching
adult size; this habitat segregation is probably related,
in part, to such factors as competition with adults and
vulnerability to predators. During postembryonic development, food conversion is expressed in maximum
growth. Adulthood represents a conservative phase in
which energy is shunted from rapid growth toward a
periodic maximization of egg output among several cohorts per year. Rapid growth and shorter lifespan are
reported to be more common in crayfishes in lower latitudes (Momot, 1984; Hazlett and Rittschof, 1985);
however, many “northern” species demonstrate these
same characteristics.
Unlike the cambarids, astacid males do not exhibit
sexual dimorphism. Copulation in Pacifastacus l. trowbridgii coincides with the autumn decline of water temperature and is soon followed by oviposition (late October and early November) (Fig. 17). Females carry
90 – 251 eggs (number directly proportional to mass)
on their abdomens, and hatching generally occurs after
7 – 8 months, during April to June. After the second
molt, the young leave the mother as 3- or 4-week old
juveniles. During the first year, individuals may undergo as many as 11 molts and nearly triple in length.
Some individuals live for eight years, and a few become
sexually mature at age three; the majority, however, become part of the breeding population during the fall of
their fourth year. This certainly follows the trend suggested by Momot (1984) that crayfishes occurring at
higher latitudes and in colder environments usually live
longer and mature later. The smallest ovigerous female
observed by Mason (1974) was 60 mm total length and
the largest 102 mm. This suggests that female astacids
lay eggs more than once (iteroparous) and, indeed, may
spawn as many as three or four times (Fig. 17). Refer
to McGriff (1983a, b) for life-history data for Pacifastacus l. leniusculus.
The life history of cambarid crayfishes (Fig. 18) is
somewhat more variable and complex than is typical of
the astacid crayfishes. The sexually dimorphic males
exhibit two distinct and usually alternating body forms
(Taylor, 1985). The change from one form to another
occurs among mature males during the semiyearly
23. Decapoda
FIGURE 17
FIGURE 18
Generalized life history of astacid crayfishes (see text for explanation).
Generalized life history of cambarid crayfishes (see text for explanation).
975
976
H. H. Hobbs III
FIGURE 19 Ventral view of basal portions of left pereiopods with
ischia bearing hooks darkened (after Hobbs, 1972).
molts. The sexually competent stage (Form I) is first attained with the last juvenile molt. This more aggressive
breeding form can be distinguished by the sclerotized,
amber-colored, and lengthened condition of the terminal elements of at least one of the first pleopods. In addition, the ischial spines are more pronounced (Fig.
19), the first chelipeds are enlarged (Stein, 1975), and
the sperm ducts are full of recently formed spermatids
(Word and Hobbs, 1958). In Form II males (the nonbreeding form), however, the terminal elements of the
first pleopod are not as well differentiated (and never
corneous), the ischial hooks are shorter and weaker,
and the chelipeds are less robust.
The ovarian cycle of female cambarids is only
partly known and from only a handful of species but
endocrine regulation of ovarian development coupled
with abiotic influences such as temperature and photoperiod are certainly paramount (Kulkarni et al.,
1991; Dubé and Portelance, 1992; Daniels et al.,
1994). Kulkarni et al. (1991) reviewed the studies devoted to this process and, working with Procambarus
(Scapulicambarus) clarkii (Girard), determined that the
ovary of this species is directly responsive to both
ovary-inhibiting and ovary-stimulating hormones.
Although Form I males may be present in a population year round (e.g., in Cambarellus and Procambarus; see also life-history data for numerous species in
Hobbs, 1981), generally they are more numerous during the fall and / or spring. Hence, many species demonstrate one or two peak periods of copulation (fall
and / or spring) although mating clearly occurs throughout the year. [Mason (1970a), Ameyaw-Akumfi (1981),
Bechler (1981), Hobbs III and Jass (1988), and Kasuya
et al. (1996) provide more detailed observations and
literature review of the mating behavior in crayfishes.]
Because oviposition occurs usually in the spring or
early summer, spermatophores (sperm plug) may be
carried by the female for as long as six or seven
months. During oviposition, the female secretes glair, a
translucent, sticky, mucilaginous substance released
from “cement glands” located on the ventral side of the
abdomen. After eggs are released from the genital pore
and simultaneously fertilized, they attach to the abdominal pleopods by a short stalk. Amazingly, it is not
known how nonmotile sperm inside a hard, waxy plug
fertilize an egg mass! Once the glair hardens, the female is considered to be “in berry.” Typically, larger females of a given species carry more eggs, yet the number of eggs is highly variable within as well as between
species (epigean forms carry as many as 600 – 700
eggs). For example, Hobbs (1981, p. 415) reported
that females of Procambarus (Ortmannicus) pubescens
(Faxon) carried between 89 and 443 eggs. He cited numerous other examples of variation in egg number for
Georgia crayfishes. Corey (1987a, b) reported on the
egg number versus individual size for three species of
Orconectes as well as for Cambarus robustus. She indicated that there was a positive linear relationship between the total number of eggs carried per female (fecundity) and the size of the female. The total number
of eggs for each was:O. rusticus ranged from 75 to
351, mean 161.8; O. virilis from 29 to 195, mean
139.1; O. propinquus from 26 to 195, mean 68.5; and
C. robustus ranged from 25 to 128, mean 64.9. See
Stechey and Somers (1995) for fecundity data on O.
immunis.
Little is known about the fecundity of stygobitic
species because ovigerous females or those carrying
young have not been observed in most species. For the
few taxa studied, however, the eggs were larger (more
yolk), but less numerous (27 – 80) (Fig. 20), reflecting
the constraints of life for these K-strategists in such energy-poor systems. Noblitt and Payne (1995) compared
water, lipid, and protein content of the eggs of P. clarkii
and P. (Ortmannicus) zonangulus Hobbs and Hobbs
and found that water contributes significantly to
weight changes. They also determined that P. clarkii
FIGURE 20 Female Orconectes i. inermis in “berry,” Marengo
Cave, Crawford County, IN (note size and number of eggs).
23. Decapoda
eggs have significantly higher percentages of proteins
while P. zonangulus eggs have significantly higher percentages of lipids. Procambarus clarkii produces relatively large numbers of small eggs (among the highest
recorded for crayfishes), whereas P. zonangulus generates relatively few but larger eggs with greater size variation. P. clarkii exhibits a profligate reproductive strategy (well adapted to warm water habitats where
nutrient input is continuous) whereas P. zonangulus
demonstrates a prudential reproductive strategy, being
better adapted to cool water environments where nutrient flow is pulsed (Noblitt and Payne, 1995; Noblitt
et al., 1995).
Depending on the season of laying and the water
temperature, eggs are carried by females for 2 to 20
weeks, during which time rhythmic movements of the
pleopods effectively aerate the developing eggs. Corey
(1987a, b) noted an inverse relationship between the
number of newly released eggs and the current velocity.
Incomplete extrusion, failure to attach to pleopods,
and fertilization failure are three other factors leading
to egg loss at oviposition. Additional losses of eggs and
young while being carried on the female result from
abrasion, predation, high current velocity, low food
availability, high density of females, and insufficient
cover for females (see also Figler et al., 1995). After
hatching, the first juvenile instar attaches to the mother
with hooked chelae and a telson “thread” anchored by
specialized hammate setae (Price and Payne, 1984).
The cephalothorax of this instar is disproportionately
large, with immense eyes, a yolk-laden carapace, and
an incompletely developed abdomen. Generally after
2 – 7 days of attachment, juveniles molt and grow to a
form more closely approximating the adult appearance.
This instar loses its posterior connection to the mother
and uses only chelae to cling to her abdomen. An additional molt usually occurs within 4 – 12 days, producing a large, third instar crayfish. Although this juvenile
hangs onto the mother with its chelae and pereiopods,
it may leave the parental pleopods intermittently; subsequent instars are free living. [Refer to Price and
Payne (1984) for more details of postembryonic ontogeny and to Mason (1970b) for maternal – offspring
behavior.] The adult female commonly molts 2 – 3
weeks after all young have left. By fall, 6 – 10 molts
have occurred in the developing juvenile, resulting in a
considerably larger and often sexually mature adult.
Mating among mature yearlings frequently takes place;
however, many individuals do not become sexually active until the following late summer or fall. Most adults
can live 2.5 – 3 years, and the majority breed more than
once. The longest life span reported for noncave
dwelling cambarids is 6 or 7 years for the burrowing
crayfish Procambarus (Girardiella) h. hagenianus
977
(Faxon) in eastern Mississippi and western Alabama
(Lyle, 1938). Longevity in stygobitic crayfishes has
been demonstrated to be considerably greater than for
epigean forms (Cooper and Cooper, 1978; Hobbs III,
1980; Streever, 1996).
C. Ecological Interactions
1. Functional Role in the Ecosystem
Crayfishes have successfully invaded a wide variety
of habitats and play important roles in processing organic matter and in the transformation and flow of energy. They are the largest and longest lived crustaceans
in freshwater ecosystems of North America and process
spatially and temporally large quantities of organic
matter (see Huryn and Wallace, 1987) in most lotic
and lentic systems. Although they can function as food
specialists, most are opportunistic generalists, feeding
virtually at all trophic levels (polytrophic) and serve as
efficient shredders and macrograzers (see Huryn and
Wallace, 1987; France and Welbourn, 1992). Shrimps,
on the other hand, are generally restricted to large,
sluggish backwaters of rivers where aquatic vegetation
(cover) is dense. They function primarily as grazers
and, like crayfishes, shrimps are eaten by a myriad of
predators (Hobbs III, 1993). Indeed, crayfishes are almost the sole source of food for some predators, such
as the striped swamp snake, Regina alleni ((Franz,
1977; Fotenot et al., 1993). Naturally, shrimps do not
play as important a role as do crayfishes in energy flow
since they are much more limited ecologically (largely
restricted to slowly moving streams or lakes and ponds
with dense vegetation). For additional information on
food and feeding by these decapods, see below.
Clearly crayfishes are important ecological constituents in numerous freshwater ecosystems yet few
studies have defined the habitat requirements for these
organisms. Taylor (1983) determined that the distribution of Procambarus (Pennides) spiculifer (LeConte)
was influenced by water depth. Rabeni (1985) ascertained that the sympatric species 0. luteus and 0. punctimanus in south-central Missouri co-exist by partitioning the physical environment. 0rconectes punctimanus
tends to be more restricted in its use of habitats, occurring as YOY in shallow water with macrophyte cover.
Both YOY and adults forage on larger-sized substrates
and both life stages prefer slow current velocities. This
species maintains a size advantage over 0. luteus (larger
individuals are dominant and occupy their preferred
microhabitat). 0. luteus tolerates a wider range of environmental conditions and exploits habitats unavailable
or undesirable to its congener. Additionally, Rabeni
(1992) determined that these crayfishes are the most
energetically important component in the diets of rock
978
H. H. Hobbs III
bass and small-mouth bass and that predation accounts
for a significant percentage of total mortality in these
crayfishes. Eversole and Foltz (1993) demonstrated that
abundance and standing crop biomass of Cambarus
(Puncticambarus) chaugaensis Prins and Hobbs and
Cambarus (Depressicambarus) latimanus (LeConte)
were altered by substrate type, velocity, cover, and water depth. For Procambarus (Leconticambarus) alleni
(Faxon) occupying the freshwater marsh habitat mosaic of southern Florida, differences in relative risk of
predation, food availability, or a combination of these
factors are probably influencing variances in habitat
occupation (Jordan et al., 1996).
Symbiotic and parasitic relationships involving
crayfishes and shrimps have been summarized by Johnson (1977), Beck (1980), France and Graham (1985),
Lichtwardt (1986), Keller (1992), Hobbs and Peters
(1993), Holt and Opell (1993), Font (1994), and
Gelder (1996). These decapods serve as intermediate or
definitive hosts and / or substrates to a wide variety of
bacteria, algae, protozoa, fungi, worms, and crustaceans. For instance, the metacercariae of the lung
fluke (Paragonimus spp.) invade crayfishes and are
transmitted to another host when the crustacean is
eaten. Paragonimus westermani is an important parasite of humans in Asia; fortunately for North Americans, few human infestations have been noted for P.
kellicoti. Ecdysis provides periodic relief from the
buildup of various ectocommensals.
Crayfishes serve as a food resource as well as a pest
for humans. A multimillion dollar business centers
around “crawfish” production in Louisiana and other
parts of the world (Huner and Barr, 1984; Avault,
1993). In addition, certain crayfishes are economically
important as local seasonal nuisances for farming. Burrowing species can be extremely abundant:Hobbs and
Whiteman (1991) reported that Fallicambarus (Fallicambarus) devastator Hobbs and Whiteman (Fig. 21)
FIGURE 21
Form I male (breeding stage) of Fallicambarus devastator, a primary burrowing crayfish from Texas.
was responsible for constructing immense burrows
with as many as 62,676 mounds / ha in the prairie section of eastern Texas! Farm equipment can obviously
be damaged, crops destroyed, and earthen dams can be
weakened by the activities of these fossorial decapods.
2. Foraging Relationships
Shrimps are somewhat opportunistic feeders, foraging on a variety of plant and animal material. The
shrimp Palaemonetes paludosus feeds heavily on epiphytic algae on vascular plants. Grazing on diatoms
(Fragilaria, Navicula, Stephanodiscus, Gomphonema,
Synedra, and Cymbella) appears to be the primary
method of food intake for populations in Hillsboro
County, Florida (Beck and Cowell, 1976); however,
they also forage for immature insects.
Barr and Kuehne (1971) reported that the stygobitic shrimp Palaemonias ganteri (Mammoth Cave, Kentucky) strained and filtered pool sediments with mouth
parts, suggesting that its food consisted of microorganisms in the silt (sediments contained protozoa; Barr,
1968). However, Lisowski (1983) observed this shrimp
exploiting another resource; an individual was observed
foraging upside-down on the surface film of a pool apparently feeding on floating material. This foraging behavior was similarly noted by Cooper (1975) for Palaemonias alabamae in Shelta Cave, Alabama, (he also
observed the sediment-straining behavior as has this author in Bobcat Cave, Madison County, Alabama).
Crayfishes play a polytrophic role in most ecosystems and, as such, contribute much to the complexities
of energy flow in aquatic systems (see above; see also
Lorman and Magnuson, 1978; Momot et al., 1978;
Momot, 1984; Hobbs III, 1993). By feeding directly on
carrion and organic debris of aquatic and terrestrial
origin, these two decapods contribute to the processing
of organic matter in aquatic ecosystems. They serve in
part as decomposers by breaking down particulate
matter, altering the chemical composition of detritus,
and releasing fecal material back into the system. Budd
et al. (1979) demonstrated that juvenile Cambarus robustus and Orconectes propinquus function as collectors by filter feeding on algae. In Missouri, YOY populations of O. (Procericambarus) punctimanus (Creaser)
possessed a greater percentage of plant detritus, O.
(Procericambarus) luteus (Creaser) had a higher percentage of filamentous algae, and both species had a
higher percentage of plant rather than animal matter in
their stomachs (Rabeni et al., 1995). Lorman (1980)
and Lorman and Magnuson (1978) showed that males
of Orconectes rusticus in Wisconsin are more carnivorous than females yet are less carnivorous than juveniles. Others (e.g., Momot et al., 1978) found that all
age classes tend to utilize vegetation as a food source.
23. Decapoda
They are capable of switching roles from herbivore / carnivore to scavenger / detritivore merely in response to food availability. It is important to recognize
that gut contents of crayfishes typically reflect availability of food type much more readily than they indicate
preference for a specific food item. Food materials constituting the majority of gut contents are not necessarily what crayfishes prefer to consume. Although individuals of some species and sizes do forage selectively,
many species are quite opportunistic and catholic in
their diets. For example, Horns and Magnuson (1981)
found that crayfishes consume significant numbers of
Lake Trout eggs in northern Wisconsin when this seasonally restricted food item is available. Savino and
Miller (1991) determined that significant impact on
lake trout egg survival would occur only in lake trout
spawning grounds with relatively low egg densities
where there was a large crayfish population, and where
cobble or large rock substrate was available. Juvenile
crayfishes are somewhat limited to detritivory and herbivory, as they filter suspended particulates and grasp
coarse particles. Adults actively prey and graze on
larger items, yet they too scrape microbes from hard
substrates and shred vegetation. The total litter processing by these crustacea was estimated to range from
4 – 6% of the annual litter input in a West Virginia
headwater stream (Huryn and Wallace, 1987). Because
of this flexibility, crayfishes can maintain large population densities despite fluctuations in food resources. Although crayfishes forage nearly continuously, the peaks
of this behavior occur shortly after sunset and during
the night. In addition, for example, this activity for
Procambarus clarkii decreased as water temperatures
began to cool to around 14°C in Oklahoma (Covich,
1978). In contrast, Price (1981), working in Arkansas,
found that both diversity in the diet and length of foraging increased for 0. (Procericambarus) neglectus
chaenodactylus Williams as food abundance decreased
during the winter months. Another factor significantly
affecting foraging is the interaction of sympatric
species, since dominance could be critical in determining both foraging and reproductive successes.
Crayfishes feed readily on macrophytes, and they
can significantly reduce species richness and biomass of
aquatic vegetation by grazing selectively on plant
species (see Lorman and Magnuson, 1978; Lodge and
Lorman, 1987; Feminella and Resh, 1989; Lodge,
1991; Creed, 1994; Lodge et al., 1994; Momot, 1995;
Nystrom and Strand, 1996). By moving through the
shallow littoral zone of lakes and cutting stems near
the bottom, they can wreak havoc even though they actually ingest very little of the vascular plant “harvest”
(an example of consumptive and nonconsumptive destruction). In contrast, Saffran and Barton (1993)
979
found that O. propinquus in Georgian Bay (Ontario,
Canada) had little impact on vascular plants, but that
they influenced primary productivity of the littoral
zone by the consumption of periphyton, charophytes,
and invertebrates.
Crayfishes, as predators (Lorman and Magnuson,
1978; Covich et al., 1981; Covich, 1982; Lodge et al.,
1987; Hobbs III, 1993; Lodge and Hill, 1994), apparently attempt to maximize energy gain with minimal
energy expenditure. For instance, rates of predation by
Procambarus clarkii on thin-shelled snails (Physa sp.)
were higher than those on thicker-shelled Helisoma sp.
(Covich, 1978). Yet, MacIsaac (1994) showed that O.
propinquus attacked medium and large zebra mussels
(Dreissena polymorpha) more often than small individuals but that predation was limited primarily to smalland medium-sized mussels. Also, O. rusticus damaged
more snails and had a greater weight-specific ingestion
rate on snails than did its congeners in Wisconsin
(Olsen et al., 1991). Hazlett (1994a) found that crayfishes feeding on zebra mussels did not respond to food
odors unless they had prior experiences with that protein source. He also determined that food odors are not
produced in sufficient quantities to stimulate feeding by
experienced individuals unless the food has been degraded by microbial activity. Results of other studies
suggest that crayfish predation reduces snail populations in lakes and streams (e.g., Lodge and Lorman,
1987; Hanson et al., 1990; Weber and Lodge, 1990;
Olsen et al., 1991; Lodge et al., 1994; Perry et al.,
1997) and also induces shifts in snail life history characteristics (Crowl and Covich, 1990). Clearly these
crustaceans have ramifying impacts on the structure
and dynamics of aquatic communities (e.g., Hanson et
al., 1990). That is, crayfishes prey upon organisms occupying various trophic levels and compete for food
and other resources at the intra- and interspecific levels.
Thus they are positioned high enough in the trophic
structure to influence numerous interactions among
several subsets of the community foodwebs (Rabeni et
al., 1995 found that Orconectes spp. accounted for
more than half of all consumption by the invertebrate
community of Missouri Ozark streams).
Foraging behavior is altered in the presence of a
predator. Stein (1977) showed that the susceptibilities
of the crayfish 0. propinquus to fish predators reflected
the size and condition of the potential prey. Life stages
with the greatest reproductive potential were those
least vulnerable to small-mouth bass predation (berried
females and Form I males). Young-of-the-year are particularly susceptible to predation; yet they, like other
size classes, must balance predation risk with the benefits of foraging. Thus, juveniles, Form II males, females,
and recently molted individuals modify their behaviors
980
H. H. Hobbs III
(reducing activity) and microdistribution (seeking substrates affording maximum protection) to minimize
risk of predation. The degree of behavioral response
appears to correlate positively with vulnerability.
The importance of top-down forces on community
composition and productivity (e.g., the trophic cascade
model — see Carpenter and Kitchell, 1992) has received
much attention (e.g., Power, 1992; Rabeni, 1992;
Strong, 1992; Carpenter and Kitchell, 1993; Lodge et
al., 1994). It is apparent that in lakes, the presence of
predatory fishes may decrease crayfish impact on lower
trophic levels by reducing crayfish abundance (Lorman
and Magnuson, 1978; Covich, 1982; Lodge et al.,
1987; DiDonato and Lodge, 1993; Garvey et al.,
1994), by nonconsumptively increasing crayfish mortality (Hill and Lodge 1995), and by reducing foraging
activity by surviving crayfish (Hill and Lodge, 1994).
Hill and Lodge (1995) further suggest that a significant
component of the impact of top predators on food
webs may be the result of sublethal effects.
Although several studies have determined the
thresholds of detection by crayfishes for various stimulatory substances (e.g., Hatt, 1984; Tierney and Atema,
1986; Hazlett, 1990; Ciruna et al., 1995), little effort
has been made to examine the role or influence of
chemical stimuli and chemoreception on the foraging
behavior of crayfishes (see Blake and Hart, 1993; Willman et al., 1994). Studies of the hierarchial structure of
food search and feeding in the spiny lobster (e.g., Zimmer-Faust and Case, 1983) suggest that chemosensoryinduced foraging in crayfishes may also be geared primarily to obtaining nearby rather than distant food
sources. Johannes and Webb (1970) have demonstrated
that several species of invertebrates release free amino
acids into the environment, and Rittschof (1980)
showed that amino acids diffuse rapidly from carrion.
Thus, amino acids are ubiquitous in aquatic ecosystems
and are readily available for use as chemical feeding
cues for crayfishes. Of particular note, Hatt (1984)
demonstrated that amino acid receptors are present on
the pereiopods and antennules of crayfishes. Tierney
and Atema (1988) suggested that the responsiveness of
Orconectes rusticus and 0. virilis to a variety of chemicals (e.g., amino acids) may reflect the omnivorous foraging habits of these crustaceans.
Stygobitic crayfishes and shrimps live in heterotrophic environments where the primary source of
energy is from the surface in the form of allochthonous
materials carried or washed into the cave, such as vegetative debris and bat guano. In addition, the stygobitic
crayfish, Troglocambarus maclanei, also is presumed to
be a filter feeder; however, cannibalism by this obligate
cave dweller has been noted (Hobbs et al.,1977). In
such resource-depressed systems, these crustaceans tend
to congregate in regions of the cave where food is spatially or temporally abundant, and thus are commonly
observed in pooled areas of streams among accumulated vegetation. Crayfish densities are particularly high
in these microhabitats, and these crustaceans often can
be observed foraging over the substrate and feeding on
microbe-encrusted debris. Their particularly enhanced
foraging efficiency reflects their improved sensory perception, reduced random movements, diminished body
size, and increased metabolic efficiencies.
Foraging behavior of burrowing crayfishes is variable; but generally on warm, humid nights, individuals
leave their burrows presumably in search of food (vegetative material). A peculiar phenomenon is noted regularly in Louisiana in which large numbers of crayfishes
(primarily males) leave their marsh habitat, move over
land, and die during the fall. They are generally in poor
physical condition, and the cause of this mass “deathmigration” is unknown (Penn, 1943, p. 15). See the
following references for additional information concerning burrowing crayfishes: Penn (1943), Hobbs
(1981), Hobbs III and Jass (1988), and Hobbs and
Whiteman (1991).
3. Behavioral Ecology
The behavioral and distributional responses of organisms to features of their physical and biotic environment is encompassed within the discipline of behavioral ecology. Behavioral responses to drought
(burrowing), orientation to current (positive rheotaxis),
light (negative phototrophism), reaction to cover, and
substrate preference (cobble – pebble), are examples of
responses by decapods to the physical environment (see
Gore and Bryant, 1990). These factors, to a large extent, determine the nature of selected habitats. Habitat
use is influenced by food, feeding rate, predators, competitors, and other sources of mortality (see Mather
and Stein, 1993). Aggressive behavior, maternal-offspring relationships, and behavioral mechanisms resulting in reproductive isolation of a shrimp or crayfish
species are representative patterns related to the biotic
features of the environment.
a. Feeding See Section III.C. 2.
b. Burrowing Atkinson and Taylor (1968) suggest
that burrowing is a strategy for minimizing adverse extremes of humidity and temperature and probably for
protection against predators; this behavior is certainly
well developed among the cambarids. Hobbs (1981)
recognized three categories of burrowing crayfishes.
“Primary burrowers” (e.g., Fallicambarus devastator,
Fig. 21) spend most of their lives in burrows, occasionally exiting to forage or to mate. Their burrows consist
23. Decapoda
981
FIGURE 22
Generalized crayfish burrows:(a, d) those of primary burrowers; (b) that of secondary burrower;
and (c, e) those of tertiary burrowers (after Hobbs, 1981, p. 32; see also Hobbs and Whiteman, 1991, Fig. 2).
of complex tunnels that are often quite removed from
open bodies of water (Fig. 22a, d) and some species
(e.g., Fallicambarus (Creaserinus) gordoni Fitzpatrick)
are found only in pitcher-plant bogs (Johnston and
Figiel, 1997). Crayfishes tend to be attracted out of
their burrows by low light intensities, and this may be
considered as an initial response that ultimately gives
rise to more complex behavior patterns involving food
searching, reproduction, and other social interactions.
Higher levels of light initiate a withdrawal response,
one that protects them from predators (see Fernándezde-Miguel and Aréchiga 1992).
“Secondary burrowers” [e.g., Fallicambarus
(Creaserinus) fodiens (Cottle) — see Norrocky 1991]
spend much of their life history in tunnels but frequent
open water during rainy seasons (Fig. 22b). Their burrows usually feature a single subvertical tube that either slopes gently or descends in an irregular spiral
(Hobbs, 1981, p. 32). “Tertiary burrowers” (e.g., Procambarus a. acutus) live in open waters and move into
burrows only at specific periods (Fig. 22c, e). For example, they burrow to brood eggs, to move below the
frost line during winter, and to avoid desiccation.
Burrows also are refugia and can be defended, and juveniles can exhibit communal living within them (see
Hasiotis, 1995). Nonburrowing crayfishes will occasionally construct burrows when surface waters disappear (Berrill and Chenoweth, 1982) or burrow into the
hyporheos of streams. For further details concerning
the burrowing behavior of crayfishes, see Grow (1981)
and Rogers and Huner (1985).
c. Water Currents Water currents certainly influence life processes of lotic organisms and bar many
species from rapidly flowing streams. Maude and
Williams (1983) demonstrated that when crayfishes are
exposed to increases in current velocity, they alter their
body posture to counteract the effects of drag and to
maintain position. Not only is this a behavioral adaptation resulting in certain species optimally occurring in
fast-flowing streams, but it may very well play a role in
the widespread distribution of species as well as in the
expansion of exotics, such as 0. rusticus, into a variety
of habitats following their introduction.
d. pH Many temperate crayfishes inhabit shallow
littoral waters during early spring. In some regions, this
time period coincides with the spring pulses of acidic
water from melting snow. Because crayfishes are capable of extensive movement and can avoid streams acidified by industrial sulfuric acid or mine drainage (pH
4.0 – 4.5; France, 1985b), they should be able to elude
environments acidified by spring melt pulses. France
(1985b) determined that Orconectes virilis can avoid
lethally acidic waters, implying that during the spring
melt, they could escape mass mortality by seeking
refuge in the profundal zone of lakes. Yet, no avoidance was demonstrated to lake water with pH values as
low as 5.6, levels that are reported to cause serious reproductive difficulties [from decreased aeration of eggs,
leading to fungal infections and egg mortality (France,
1985a)]. The failure of this crayfish to avoid sublethal
acid waters could, therefore, interfere with recruitment;
982
H. H. Hobbs III
for this reason, France suggests that the prolonged survival of 0. virilis in waters receiving acid runoff is
doubtful. This may be the case for other crustaceans
living in poorly buffered systems but, surprisingly
enough, some crayfishes [e.g., Procambarus (Ortmannicus) fallax (Hagen) and P. paeninsulanus] reach maximal densities in acid waters. Yet, Tierney and Atema
(1986) experimentally showed that acid exposure inhibits behavioral responses (specifically feeding and
grooming; see also Uiska et al., 1994); this could cause
acid-stressed populations to decline at pH levels well
above those at which lethal physiological effects become apparent.
e. Shelter and Aggression Obviously important aspects of the ecology of crayfishes and shrimps are their
intra- and interspecific behavioral responses. Although
they are nonterritorial, they are aggressive and rely on
agonism to procure shelter, food, and mates (Stein,
1975). Most crayfishes tend to be solitary and occupy
and defend crevices in a coarse substrate, both of
which are of survival value. Crevices not only provide
shelter from pounding shore waves or from high
stream velocities, but they also furnish a refuge that
can be defended against predators and competitors (see
Blank and Figler, 1996). Considering the generalistic
feeding habits of crayfishes and the apparent absence of
food limitation in most systems, the principal resource
bottleneck may be crevice availability. Hobbs III (1976)
demonstrated a positive relationship between crayfish
abundance and availability and size of cobble substrates. Caine (1978) observed that competition among
Florida epigean crayfishes is primarily for shelter, and
individuals unable to obtain cover succumb to predators. A specific home range has been demonstrated for
a number of species (e.g., Hazlett et al., 1974; Hobbs
III, 1980), and dispersion and dispersal often appear
related not to population density but to water depth,
number of crevices, predators, temperature, etc. Experiments examining intraspecific aggression of resident
maternal (carrying eggs and / or hatchlings) P. clarkii
against intrusions by Form I males and nonmaternal female conspecifics showed that maternal residents won
a significantly higher proportion of encounters with either sex, thus demonstrating maternal aggression in
crayfishes (Figler et al., 1995). Kershner and Lodge
(1995) showed that O. rusticus’ use of cobble habitat
in northern Wisconsin lakes was significantly and positively correlated with lakewide predator density, reflecting the potential importance of predation risk in
establishing crayfish distribution patterns. They suggested that the availability of cobble habitat may influence the success of the invasions of this crayfish. Additionally, crevices serves as a feature, which, by further
partitioning the environment, permits a greater species
richness in those habitats.
Stein (1975) indicated that the chelipeds of O.
propinquus make up 40% of their adult total dry
weight. These immense pereiopods are used in sexual
encounters with females, for procuring food, and in agonistic confrontations. Rutherford et al. (1995) demonstrated that chela length (but not symmetry) is positively related to fighting ability of O. rusticus.
Levenach and Hazlett (1996) found that if the display
function of chelae is impaired then the crayfish is prevented from competing successfully with conspecifics
(O. virilis), yet impairing the mechanical function of
chelae does not reduce the crayfishes’ competitive ability. Since crayfishes can autotomize their chelae, Figiel
and Miller (1995) investigated the influence of chela
autotomy on growth and survival of P. clarkii. They
showed that as population density increased, the frequency of chela loss increased and that injured individuals demonstrated reduced growth and survival rates.
Hayes (1975) demonstrated eight innate components of social interaction in the primary burrowing
crayfish, Procambarus (Girardiella) gracilis (Bundy):
alert, approach, threat, combat, submission, avoidance,
escape, and courtship. These also have been observed
in other species and Ameyaw-Akumfi (1979) summarized the sequence of behaviors demonstrated by interacting crayfishes during aggressive encounters (Fig. 23).
He proposed that movement of gnathal appendages
and pereiopods as well as repeated antennal waving occurs prior to the termination of an agonistic encounter
and this behavior may in fact constitute an appeasement display. He further stated that “the use of appeasement signals in mating behavior is, probably, a
capitalization by the male as this behavior appears to
reduce aggressive tendencies” (Ameyaw-Akumfi 1979,
p. 41). Copulatory behavior (reviewed in Bechler,
1981) is generally seasonal with the pair assuming several positions, the male mounting the female most commonly (Fig. 24). Mating generally lasts from 10 min to
10 h (e.g., Andrews, 1895; Mason, 1970a). After
oviposition, egg fanning, and hatching, maternal – offspring behavior is mediated by pheromones released
into the water by the female. These cues are speciesspecific but not brood-specific, since juveniles are attracted to conspecific brooding females. Also, Peck
(1985) showed that social interaction is an important
factor affecting temperature selection by 0. virilis. Individuals demonstrated behavioral mechanisms, not only
to find favorable temperature ranges but also to segregate according to social rank.
f. Predation Avoidance The presence of predators
elicits various behavioral response in crayfishes. Young-
23. Decapoda
983
FIGURE 23
Agonistic behavior between two male Orconectes propinquus in the “body up”
position; crayfish on right performing “antenna tap” (Bruski and Dunham, 1987) (photograph
courtesy of Ann Jane Tierney).
of-the-year are particularly susceptible to predation
and must balance predation risks with foraging benefits
(Butler and Stein, 1985). Stein and Magnuson (1976)
and Stein (1977) showed that in the presence of smallmouth bass (Micropterus dolomieui), 0. propinquus
foraged less, adopted defensive postures, and sought
protective substrates. Ideally, crayfishes use crypsis to
avoid detection by predators. However, when discovered, crayfishes typically backswim seeking shelter in
vegetation, beneath rocks or debris, or in burrows. If
FIGURE 24 Copulating pair of Orconectes propinquus, Form I male on top (photograph courtesy
of Ann Jane Tierney).
984
H. H. Hobbs III
escape is not possible, individuals assume and intensify
a species specific defensive posture, the predator response posture (PRP). Hayes (1977) described PRP of
crayfishes and discussed the evolutionary significance
of these behavioral patterns in three species of the
genus Procambarus. The crayfishes studied have similar
lateral merus displays which result in a lateral presentation of the chelae, allowing for defensive coverage of
the entire carapace and for quick flight by tail-flipping.
Also, elongated chelae of these species enable them to
sense tactilely the approach of a predator and to keep it
at a greater distance. The PRP of the terrestrial (burrowing) species P. gracilis appears to have been modified for threat from terrestrial predators with attack
coming primarily from above. Chelae are held shieldlike in front of the carapace, and the tail is folded
tightly beneath the body, allowing for rapid reorientation to the attacks of a shifting predator. This posture
frees the tail from use as an escape organ on land. Stein
and Magnuson (1976) also suggested that, in addition
to their effects on the distribution and behavior of
crayfishes, predators have an influence beyond simple
predator – prey interactions because of their impact on
several trophic levels.
4. Population Regulation
The population dynamics of shrimps and crayfishes
are not well studied except for a few species. Traditionally, the four major features of any demographic model
are summarized as natality, mortality, immigration, and
emigration. Each of these clearly affects the size and
growth of natural populations and all are under the
constraints of density-dependent (D-D) and / or densityindependent (D-I) factors. It should be emphasized,
however, that conventional D-D and D-I regulatory
factors, while present, are too often superceded in importance by the effects on density of human activities
(Hobbs and Hall, 1974). Rather than a single factor, it
should be emphasized that any of these elements may
interact strongly in affecting shrimp and crayfish community composition, population sizes, and production.
a. Shrimps A 1-year life history is characteristic
of epigean shrimps, whereas cave forms probably live
longer. Food resources tend not to be limiting for
epigean shrimps, but fish predation can have significant
impact on their densities (Nielsen and Reynolds, 1977).
The same investigators noted that fluctuating water
conditions (seasonal drying and extended high water)
also elicit population changes. Fecundity appears to be
related to size (length) of female and to time of year.
In cave environments of the southeast, amblyopsid
fishes (and crayfishes) are predators of stygobitic
shrimps and crayfishes. In these food-limited systems,
where shrimp and some crayfish populations are very
small, loss of ovigerous females (and thus loss of recruitment of young) can be very significant to immediate and particularly long-term stability of population
densities. Lisowski (1983) attributed the extreme decrease in density of Palaemonias ganteri from baselevel streams in the Mammoth Cave System, Kentucky,
to pollution of local groundwater. The sources of contaminants were human sewage and water from the
Green River, which had received inputs of oil brines
and hydrocarbons during the 1960s. He also indicated
that modifications of habitat and flooding regimes
caused by dams within the Green River drainage adversely affected shrimp by reducing the reproductive
success and by increasing predation pressure on them.
b. Crayfishes Lodge and Hill (1994) reviewed the
variables regulating population size and the importance
of both D-D and D-I factors for some cool-water crayfishes. They summarized the D-I parameters as temperature, calcium concentration, pH, dissolved oxygen,
salinity, habitat (substrate particle size and food availability), and water velocity and indicated that the D-D
factors are diet, competition (intra- and interspecific),
predation, molting, and ectosymbionts and disease.
Momot and others, in a variety of population studies
(Momot and Gowing, 1977a, b; Gowing and Momot,
1979; France, 1985a), have shown that various populations of 0. virilis are regulated by several D-D and DI factors, including cannibalism, natural mortality at
molting (physiological and mechanical problems), predation by snakes (Godley et al., 1984), fishes (e.g.,
trout, bass) and some insects (e.g., dragonfly) on yearlings, predation on adults by various terrestrial vertebrates (Lodge and Hill, 1994), flooding, temperature,
pH (Davies, 1984; Siewert and Buck, 1991; France and
Collins, 1993), seiches (Emery, 1970), and other factors. Crayfishes are generally characterized by high
juvenile mortality and reduced adult mortality (concave
survivorship curve). It has been suggested (Momot and
Gowing, 1977b) that overall, most changes in biomass
and productivity of these populations result from internal density-dependent processes operating on fecundity
and mortality rates rather than on growth. In several
Michigan lakes during most years studied by Momot,
the breeding success of 0. virilis depended mainly on
the survival and subsequent fecundity of the 1.5-year
age group females. At 2.5 years, females are considered
an important strategic reserve.
Where natality, mortality, emigration, and immigration rates vary, population sizes will fluctuate, especially where birth recruitment is inadequate. As populations expand, they face potential resource depletion
and the possibility of attracting more predators — both
23. Decapoda
factors that can limit population size. Predation pressure may cause prey to modify their behavior (e.g., increased burrowing and chelae displays, suppressed foraging, and reduced overall activity). In turn, these
antipredatory behavioral patterns act as selective (coevolutionary) forces on the predator, thus operating to
modify predator strategies. Stein and Magnuson (1976)
and Stein (1977, 1979) have demonstrated that juvenile
O. propinquus are the preferred crayfish prey of smallmouth bass and, as such, are rarely found on open,
sandy substrates. Adult crayfish (least preferred prey)
are abundant in these areas, demonstrating both that
preferred prey make distributional shifts to protective
habitats and that use of space by nonpreferred prey is
regulated much less by predators.
Inter- and intraspecific competition for habitat
(refuge) and nutrients (food) affects crayfish populations in a variety of ways. When food or shelter are
limiting crayfishes interact agressively, suggesting that
interference competition may be common (Capelli and
Munjal, 1982; Capelli and Hamilton, 1984). Bovbjerg
(1956) noted that larger size entrusts a competitive advantage; and during intraspecific competitive encounters, male crayfish usually dominate female crayfish.
Also, others (e.g., Flynn and Hobbs III, 1984; Hazlett
et al., 1992; Lodge and Hill, 1994) have suggested that
interspecific competition for habitat and food results in
competitive exclusion.
Length – frequency distributions have traditionally
been used by investigators to show age and growth of
individuals in populations (see France et al., 1991 for
conditions required to produce reliable results).
Growth of crayfishes is related directly to the number
of molts during a growing season and the growth increment per molt; growth is stepwise rather than continuous. Average growth increments range from 0.41 mm
for the dwarf crayfish Faxonella clypeata (Hay)(Black,
1958) to 2.6 mm for Procambarus clarkii (Penn, 1943).
The difference in yearly size increment between males
and females is due primarily to the frequency of molting and not necessarily to the growth increment at each
molt (Hazlett and Rittschof, 1985). However, the increment for individuals living in optimum conditions may
well have as much impact as does the number of molts.
Female growth is depressed, probably in response to
the effects of carrying eggs and young, resulting overall
in fewer molts.
Conditions that promote rapid juvenile growth
rather than adjustments in egg production per se modify any stock recruitment interaction. In more northern
climates, the temperature of the water in the shallows,
especially in “nursery areas,” is an important determinant of the molting rate in juveniles. Flint (1975) found
that a coastal stream population of Pacifastacus lenius-
985
culus had a slightly faster overall growth rate than a
subalpine lake population; he ascribed this to higher
water temperatures and a longer growing season. Because crayfishes molt more frequently as juveniles, any
parameter reducing juvenile molt frequency also reduces growth rate; this, in turn, may determine size at
maturity, although there is no absolute size at which a
crayfish becomes mature. Because fecundity is a function of female size, populations with smaller females
should have lower recruitment of YOY. Obviously, a
cohort that grows faster and reaches a larger overall
size will have a better chance of ensuring the replacement of its population. Huryn and Wallace (1987)
demonstrated that long life span (five years) and slow
growth of Cambarus bartonii in a West Virginia headwater stream resulted in low production (see also Roell
and Orth, 1992); biomass exceeded production and
transpired in a production / biomass ratio (P / B) of 0.58,
a value similar to those reported from other temperate
crayfish populations (e.g., Momot, 1984). Working
with long-term data sets for Orconectes virilis in two
Ontario, Canada lakes, Momot and Hauta (1995) suggested that crayfishes can have cohort P / B ratios ranging from 1 to 8 and that fluctuations in growth and
mortality can alter these ratios significantly.
Crayfishes commonly establish sole occupancy of
protective crevices in coarse substrates (when available)
which they defend agonistically. A study conducted by
Aiken (1968) in Alberta found that both male and female Orconectes virilis move to deeper water in late
summer, females preceding males. Severe winter conditions would eliminate a population if it remained in the
littoral zone where water freezes to the bottom. Rather
than burrow to avoid ice, individuals escape by migrating to deeper water. Young-of-the-year appear to have
higher mortality rates during the winter than adults. At
spring thaw, both males and females return to shallow
water. In streams, crayfishes generally burrow into the
hyporheic zone to overwinter.
Periods of low water obviously affect all members
of the aquatic community, yet crayfishes can escape
desiccation by burrowing into the gravel armor of
stream channel beds. Prior to complete loss of an
aquatic habitat, other responses by crayfishes have
been noted. Taylor (1983), studying a lotic population
of P. spiculifer during a drought in northeastern Georgia, suggested that population shifts (decrease in adult
population densities, change in reproductive timing, increase in number of juveniles) resulted from loss of preferred deep-water sites, emigration, and subsequent increase in predation of larger adults.
Population regulation among sympatric species has
received considerable attention in the last 15 – 20 years
(e.g., Lodge et al., 1986; Lodge and Hill, 1994). Many
986
H. H. Hobbs III
North American crayfishes occur sympatrically with
close relatives, yet hybridization appears to occur relatively infrequently although it may be much more common than previously recognized (see Roush, 1997).
One mechanism that may be operating to ensure reproductive isolation is chemical or visual recognition of
species during mate selection (Smith, 1988). Chemoethological isolating mechanisms could be effective for
sympatric but not allopatric species. In consequence,
hybridization of a resident and an exotic species might
be more likely than between two distinct but geographically overlapping resident species. However, morphometric analyses indicate a strong likelihood that hybridization does occur between O. rusticus and O.
propinquus in northern Wisconsin lakes where the former is an exotic. Lodge and Hill (1994) report that putative hybrids comprise up to 27% of mixed-species
populations, and laboratory crosses have shown that
viable offspring from matings of these two species are
produced but in numbers that are 60% lower than in
conspecific matings (Berrill and Arsenault, 1985). Also,
Cesaroni et al. (1992) report a narrow hybrid zone
maintained by two closely related undescribed crayfishes (Procambarus spp.) cohabiting a cave in northern
Chiapas, Mexico.
Sequential invaders (natural or introduced) can
elicit drastic changes in community structure (Hobbs
III et al., 1989), including replacement of native species
(Hill et al., 1993; Kershner and Lodge, 1995; Gamradt
and Kats, 1996). In recent years, biological invasions
have been considered such a major environmental
problem that many volumes have been devoted to their
study (see Lodge, 1993a). Of particular importance to
crayfish biology is the apparent replacement of native
crayfishes in various parts of North America (and
world, Hobbs III et al., 1989) by introductions of nonnative species (e.g., interactions of 0. virilis, 0. propinquus, and 0. rusticus in Wisconsin, Lodge; 1993b;
Capelli, 1982; Capelli and Magnuson, 1983; Page,
1985; Hobbs III and Jass, 1988; Momot, 1992;
Charlebois and Lamberti, 1996; Taylor and Redmer,
1996). Displacement mechanisms are complex, and the
mechanisms regulating shifts and / or replacements of
species are not well understood in most instances
(Flynn and Hobbs III, 1984). However, field and laboratory results indicate that reproductive interference,
growth, predation, competition, aggressive dominance,
mortality, and interspecific morphological differences
probably serve as mechanisms regulating replacement
(Capelli, 1982; Capelli and Munjal, 1982; Butler and
Stein, 1985; Lodge et al., 1986; Mather and Stein,
1993, Hill et al., 1993; Hill and Lodge, 1994; Garvey
and Stein, 1993; Garvey et al., 1994). Based on data
from various lakes in northern Wisconsin, Lodge et al.
(1986) and Lodge (1993b) stated that outcomes of
such invasions are not necessarily predictable and
Lodge et al. (1998) suggested that “best-guess” models
of invasions, short of experimental work before or during an invasion, is the best approach. This unpredictability is due to a complex suite of factors such as
reproductive success, competition for resources, diel
and year to year fluctuations in predation risk, and
parasitism. A variety of disturbances are involved in
the “final” community structure (see Olsen et al., 1991;
DiDonato and Lodge, 1993; Garvey and Stein, 1993;
Hill and Lodge, 1994).
IV. CURRENT AND FUTURE
RESEARCH PROBLEMS
Even though considerable effort has been directed
toward the study of these decapods, with particular
emphasis placed historically on taxonomy, systematics,
evolution, anatomy, and some areas of physiology, satisfying answers only to a few questions concerning
them have been obtained. Barely a handful of functional morphology studies have been conducted. Recent realization of the impact of exotic species introductions (e.g., Orconectes rusticus, Capelli, 1982;
Magnuson and Beckel, 1985; Hobbs III and Jass, 1988;
Hobbs III et al., 1989; Lodge, 1993a) has initiated numerous studies to determine the underlying mechanisms controlling crayfish species replacements and
community changes (Butler and Stein, 1985; France
and Welbourn, 1992; Mather and Stein, 1993; Garvey
et al., 1994).
Our knowledge of life histories of shrimps and crayfishes is incomplete, at best. As noted above, only about
20 species have been studied in any detail, and varying
quantities of data exist for the remaining taxa. Although
much time and energy are required for these research
projects, the need is paramount. Detailed studies of
ovarian development and oogenesis as well as testis development and production of spermatozoa are sorely
needed. Also, much is lacking in our understanding of
the dynamics of populations for virtually all species of
freshwater decapods. Best known are factors influencing
population dynamics in the crayfish 0. virilis.
The Fish and Wildlife Service of the U.S. Department of Interior periodically issues a List of Endangered
and Threatened Wildlife. Currently there are seven decapod species placed on the Endangered list, four of
which are stygobites. Palaemonias alabamae and P. ganteri are stygobitic shrimps in Alabama and Kentucky,
respectively; Cambarus (Jugicambarus) aculabrum
Hobbs and Brown is known only from two caves in
northwest Arkansas and is the “newest addition” to the
23. Decapoda
Federal List (Jacobson, 1996; Elliott, 1997; Boyd,
1997); and Cambarus (Jugicambarus) zophonastes
Hobbs and Bedinger is found in an Arkansas cave. The
crayfishes Orconectes (Crockerinus) shoupi Hobbs and
Pacifasticus fortis are restricted to streams in western
Tennessee and Shasta County, northeastern California,
respectively, and the shrimp Snycaris pacifica inhabits
streams in central California. Additionally, the Fish and
Wildlife Service listed the stygobitic shrimp, Palaemonetes cummingi Chace, as a Threatened species. The
1996 IUCN Red List of Threatened Animals (Baillie
and Groombridge, 1996) cited three atyid (2 Palaemonias, 1 Syncaris) and two palaemonid shrimps (Palaemonetes); two astacid (Pacifastacus) and 118 cambarid
(3 Camabrellus, 34 Cambarus, 2 Distocambarus, 8 Fallicambarus, 6 Hobbseus, 24 Orconectes, and 41 Procambarus) crayfishes as Threatened; and the atyid
shrimp Syncaris pasadenae is listed as Extinct.
The American Fisheries Society Endangered Species
Committee listed all crayfishes occurring in the United
States and Canada and reviewed the conservation status of all taxa (Taylor et al., 1996). There are state lists
as well (e.g., Florida — Deyrup and Franz, 1994).
Clearly, it is important that these species and their critical habitats be monitored.
The ecology of decapods inhabiting resource limited cave habitats is poorly understood (see Hobbs III,
1992; Brown et al., 1994; Streever, 1996). Further investigations should be conducted to determine the
physiological mechanisms that allow the reduction of
energy utilization in tissues of stygobites. These should
be run in conjunction with studies delineating the levels
of environmental flexibility of these forms.
Brown (1981), using electrophoretic analysis of 19
biochemical loci of six crayfishes, substantiated previous reports of low genetic variability in decapod crustaceans [(also demonstrated for crayfishes by Fuller et
al. (1989), Crandall (1993), Koppelman and Figg
(1995), and Fetzner (1996)]. She attributed the low
heterozygosity in these crayfishes from the southeastern
United States to their limited geographical distribution
in a low number of drainage systems and to their high
mobility within those watersheds — both are factors allowing crayfishes to perceive their habitats as finegrained. Nevo (1978) proposed that generalists are
more genetically variable than habitat specialists
among all major taxonomic groups and Brown, on the
basis of her findings, suggested that crayfishes should
be classified as habitat specialists. Biochemical investigations of other crayfishes occupying a variety of habitats and from different phylogenetic positions should
be undertaken to test the latter hypothesis. Also, additional allozyme studies as well as nucleotide sequence
data from the amplified mitochondrial DNA from the
987
16S ribosomal subunit may aid in determining more
precisely the relationships of currently recognized genera, subgenera, species groups, species, and subspecies
(see Fetzner, 1996; Crandall and Fitzpatrick, 1996).
Current hypotheses of taxonomic relationships may
not reflect actual phylogenetic relationships.
Recently, considerable effort has been directed toward the study of pheromones and other chemical substances released into the environment by various decapods. Much is to be learned about the importance of
chemical communication in the behavior of aquatic animals and about the impact of these substances on the
structure of communities. Basic questions remain unanswered concerning the roles of pheromones and chemical cues in population and community dynamics. Obviously, a tremendous variety of behavioral studies await
the ethologist.
Although James (1979, see his bibliography) has
shown that benthic macro invertebrates are good indicators of various forms of pollution and are useful as
test organisms for determining the presence of toxic
substances, the importance of the role of freshwater decapods has not been completely demonstrated (Vermeer, 1972; France, 1986). Recent technical advances
and syntheses have brought together existing data on
the effects and measurement of toxicity in decapods
(Hobbs and Hall, 1974; Doughtie and Rao, 1984;
Roldman and Shivers, 1987; Wright and Welbourn,
1993). Acidification due to acid precipitation and loss
of buffering capability in freshwater systems in the
northeastern United States and Canada have stimulated
research on the effects of increased acidity on organisms (France and Graham, 1985; Tierney and Atema,
1986; Wheatly et al., 1996). Crayfishes are highly sensitive to an increase in the hydrogen ion concentration
(Wood and Rogano, 1986; Bendell-Young and Harvey,
1991) and although there are inter- and intraspecific
variations in the degree of tolerance (Berrill et
al.,1985), France (1984, 1985a) has shown that Orconectes virilis juveniles are particularly sensitive. DiStefano et al. (1991) determined that adult C. b. bartonii
and C. robustus are among the most acid-tolerant
freshwater organisms they tested. This implies that if
populations of these two species would begin to be affected negatively by increasing acidification, then it
may already be too late for the recovery of most other
species in those lotic or lentic habitats (see Clancy,
1997). Additionally, studies are needed in order to determine the impact of toxic materials on spermatozoa,
ova, developing embryos, young, and individuals in
various molt stages (premolt, postmolt, molt, intermolt). For those species that regularly experience dilute
seawater in some portions of their ranges (e.g., Pacifastacus leniusculus and P. clarkii), relatively little is
988
H. H. Hobbs III
FIGURE 25
Practice of “ditching” to lower local water table significantly (note person for scale) is a
common practice (Hillsborough County, FL) with numerous negative environmental impacts.
known concerning the identity and character of short
and long-term control of the sodium pump in the crustacean gill and antennal gland (Sarver et al., 1994).
Much is yet to be discovered about the physiology of
the gut, including enzyme systems, digestion and absorption of nutrients, and metabolic fate of absorbed
nutrients.
As a result of increasing demands by humans for
energy, aquatic ecosystems undoubtedly will be subjected to considerable additional waste heat. Because
crayfishes and shrimps occupy important positions in
complex food webs of lentic and lotic ecosystems, precise data concerning temperature preferences, heat tolerances, and thermal responses (e.g., McWhinnie and
O’Connor, 1967; Claussen, 1980; Mathur et al., 1982;
Taylor, 1984) of these and other organisms will be invaluable in establishing temperature standards such
that aquatic communities will be protected.
The following is only a partial list of threats to
freshwater ecosystems. A thorough understanding of
these and other potential sources of perturbations is
paramount if we are to attempt to protect freshwater
organisms, including shrimps and crayfishes. Point and
nonpoint agricultural (e.g., pesticides, sediments) and
other sources (e.g., municipal runoff and discharge) of
pollutants; urban and residential development; heavy
recreational use of private and public lands (e.g., horse
camps); gem, feldspar, coal, and mica mining; logging
and chip mills; paper mills; industry (particularly chemical); impoundment structures (dams) and their opera-
tions; highway expansion; dewatering of stream sections (e.g., Hiwassee River), including “ditching” (Fig.
25); disturbances (chemical, physical) to hydrological
systems in karst areas; sport fishery (stocking, baitbucket introductions); pig lots; disruption of riparian
vegetation; acid rain; and catastrophic spills (highways,
trains). Sadly, this is only a brief start to a lengthy list
of potential threats to freshwater ecosystems.
Much effort is and will continue to be required in
order to recover disturbed lotic and lentic habitats and
to conserve and preserve existing natural ecosystems
(Richter et al., 1997); this includes surface and subsurface water resources. Aside from basic research conducted to recognize contamination and to treat perturbed areas, implementation of new laws, education,
and stricter enforcement of existing legislature are necessary strategies of the future. In some parts of the region treated in this book (e.g., Louisiana), shrimp and
crayfish production are important economic activities.
Considerable work has been directed toward increasing
the yields of crayfish farmers and this effort should be
continued (Avault, 1993; see also Daniels et al., 1994).
As marine shellfish populations are depleted, humans
will undoubtedly turn to alternate food sources, including crayfishes. In various countries in Europe, these
crustaceans are valued as food and extensive stocking
of exotic species (extreme caution advised!), and rehabilitation programs for native species are ongoing.
Most of the interest in North America is centered on
development of an export trade for Europe, yet there is
23. Decapoda
potential for an expanded domestic market. Those
crayfish species with known economic importance in
North America, such as the burrowers Cambarus (Lacunicambarus) diogenes Girard and Failicambarus devastator, the stream and lake-dweller Orconectes rusticus, and the tasty Procambarus clarkii, particularly
should be studied in detail.
V. COLLECTING AND REARING TECHNIQUES
Because decapods are found in such a large diversity of habitats, a variety of techniques should be used
in procuring representatives.
A. Shrimps
1. Atyidae
Collections should not be made for any of the atyid
shrimps in North American freshwaters. Populations of
the two species of Syncaris and the two cave shrimps assigned to Palaemonias are very small (two may already
be extirpated) and should not be disturbed!
2. Palaemonidae
Freshwater shrimps occupy various sluggish lotic
or lentic environments and are most readily collected
with the aid of a sturdy, long-handled, fine-meshed dip
net or a small-meshed seine. When a particular locality
is choked with vegetation or if the water is greater than
1.5 m deep, the use of fine-meshed wire traps with inverted cones baited with meat can be employed. In
clear to low turbidity waters, they may be collected at
night with the aid of a headlight and dip net (the rubycolored reflection of their eyes makes for easy location
of individuals). Shrimps can be maintained in well-aerated aquaria with considerable vegetation; the water
should be changed weekly.
B. Crayfishes: Astacidae and Cambaridae
The North American astacids are known primarily
from lakes and streams west of the Continental Divide,
and methods for collecting these crayfishes are identical
to those for the cambarid stream / lake-dwellers. In
shallow streams that have little vegetation, a smallmeshed seine (2 – 5 m long) can be most useful. Deep
sections or pooled areas of streams as well as shallow
ponds and the littoral zone of lakes may be sampled by
pulling the seine (5 – 8 m length) through the water,
taking care to ensure that the weighted margin of the
seine remains in contact with the substrates. When
electrofishing equipment is used, small-meshed seines
989
can collect any organism that is stunned and carried
downstream. Use of various wire traps is recommended
for vegetation-choked streams, swamps, marshes,
ponds, or lakes or when sampling deep waters of lakes.
Inverted-cone minnow (funnel) traps with enlarged
openings (4 – 5 cm diameter) have proven to be quite
effective. A residence time of 1 – 2 nights, using chicken
and fish scraps, liver, canned cat food, etc. as bait, is an
adequate and productive duration for this sampling
technique but this method suffers from sampling bias
(Stuecheli, 1991).
A sturdy dip net is very helpful in sampling ditches,
mud-bottomed pools, overhanging vegetation, and areas choked with vegetation. This type of net (delta net
preferred) can be used to collect individuals from rocky
substrates in lakes and streams. Probably most crayfishes burrow at least occasionally for one or more reasons. The diameter of the burrow tube and chimney
pellet size provide good indicators of the size of the
crayfish inhabiting the burrow (Hobbs III and Jass,
1988). One may examine the burrow aperture and assume that the smaller openings contain juveniles; to
sample the adult population, concentrate on burrows
with larger diameters. [This would not be an appropriate procedure for small burrowing species, such as
Cambarellus (Pandicambarus) puer Hobbs, Faxonella
clypeata, and Procambarus (Hagenides) pygmaeus
Hobbs]. On warm, rainy, humid nights, burrowing
crayfishes can be collected fairly easily. These crayfishes
come to the mouths of their burrows and often leave
them to forage for food. With the aid of a headlight,
they can occasionally be collected in large numbers (see
also Norrocky, 1984). Use of scuba equipment to observe and / or collect crayfishes in deep streams, pools,
ponds, and lakes also is a very useful technique [see
France et al. (1991) and Lamontagne and Rasmussen
(1993)].
During population studies where long-term data
are required for individual crayfish, various marking
(tagging) techniques are employed. Because these crustaceans periodically molt and thus lose their exoskeleton, external tags are not used since they interfere with
casting off the exuvium or they are lost when the exoskeleton is shed. Consequently, internal tags are recommended [see review in Hobbs III (1980); Weingartner (1977) for colored filiaments; Wiles and Guan
(1993) for microchip implants].
Stygobitic decapods generally lack pigments and
thus appear white or translucent. In subterranean
streams, use of a small aquarium net is generally
adequate to capture the readily visible organisms.
Hand-grabbing also is a fairly productive means of collecting them once they have been disturbed and are
“swimming” in open water. Wire minnow traps can be
990
H. H. Hobbs III
used if checked often. [Note: cave decapod populations
are usually small and because of reduced biotic potentials in stygobitic forms, yearly recruitment is very low.
Thus, care must be taken not to collect many (if any)
individuals from any cave locality — at any time!]
Because crayfishes are aggressive, antisocial, and
cannibalistic, rearing and maintaining them can be
problematic. On the other hand, because they are scavengers and detritus feeders, they will feed on almost
anything organic. Ideally, individuals should be maintained separately, each crayfish in a single container
(e.g., 20 8 cm stackable glass bowl) and at room
temperature. No aeration is required, provided the water does not become fouled with food, wastes, etc. The
bowls should have small gravel substrates, and the water should be sufficiently deep to cover the crayfish. If
larger containers are used, small cobbles limestone
fragments, or broken pieces of flower pots, bricks, or
PVC pipe cut to varying sizes can be added for cover
(see Alberstadt et al., 1995). Individuals should be fed
2 – 3 times per week, and the water should be changed
approximately every other week. Food can range from
various aquatic plants (Potamogeton, Elodea), scraps
of meat, and earthworms to dried cat food. Continuous
illumination will ensure a growth of algae but is not
necessary.
Shrimps can be raised in much the same manner as
described for crayfishes except more vegetation and
deeper water are required for these crustaceans. Females with eggs should definitely be kept isolated.
When young hatch, they will remain attached to the
pleopods of the female for up to several weeks. Most of
the young will then leave the mother and the female
should be separated from them to prevent parent / offspring cannibalism. As the young undergo molting and
growth, some sibling cannibalism will probably occur.
By providing cover for the molting young or by dividing them among several containers, this can be reduced
significantly.
On a much grander scale, large tanks or ponds can
be utilized for raising crayfishes or shrimps. These
should be designed for complete draining with sluice
gates or standpipes and should be long and narrow
rather than square (Figure 26). Water depth should be
maintained at about 1 – 1.5 m and food can be variable, but catfish feed or poultry starter are good. For
additional information on various aspects of crayfish
aquaculture, see Huner and Barr (1984), Avault
(1993), and Huner (1995).
Upon collection, crayfishes and shrimps should be
killed in 5% neutral formalin and kept in the solution
from 12 h to a week, depending on the size and number of individuals. Specimens should then be washed in
running tap water for several hours and transferred to
70 – 75% ethanol. If epizooites (e.g., protozoans, copepods, entocytherids, branchiobdellids) are to be saved,
FIGURE 26 Aquaculture ponds for raising crayfish (the blue color morph, Procambarus paeninsulanus) in Hillsborough County, FL.
23. Decapoda
991
the formalin in which the decapods were killed should
be filtered, the exoskeleton rinsed, and the rinse water
poured through a fine seive. Any symbionts should be
preserved in 75% ethanol and carefully labeled (Hart
and Hart, 1974). Where possible, crayfishes and
shrimps should be stored in clamp-top glass jars with
gaskets. The organisms should be placed in the jar “anterior end down” with room for adequate 70% ethanol
preservative. The specimens should be accompanied by
a 100% rag paper label on which data are clearly
printed using insoluble black ink.
VI. IDENTIFICATION
A. Preparation of Specimens
Reliable identification of freshwater shrimps can be
made only if appendages are removed, cleared, and
mounted on a microscope slide. The second pleopod of
adult males should be extirpated and then heated in a
lactic acid – chlorazol Black E stain solution [3 – 5 drops
in 1% ethanol (95%) stain solution per 20 mL of lactic
acid; a 1% solution of fast green can be substituted] at
150°C for 15 min. The translucent appendage should
be transferred to glycerine and then examined.
Crayfishes can be identified with the aid of a hand
lens or a stereoscope. First-form males are required
FIGURE 27
Caudal view of first pleopod (gonopod) of first-form
males (after Hobbs, 1972). (a) Bouchardina robinsoni Hobbs (symmetrical); and (b) Procambarus (Ortmannicus) acutissimus (Girard)
(asymmetrical); (pms, proximomesial spur).
for positive identification, and a crayfish’s left pleopod should be removed and used for working with
the key.
B. Taxonomic Key to Genera of Freshwater Decapoda
Morphological characteristics used in the following
key are illustrated in Figures 1, 2, 19, and 27 – 29.
First-form males are required to identify crayfishes at
the generic level.
la.
Rostrum and abdomen compressed laterally; third pair of pereiopods never bearing chelae ................................infraorder Caridea 2
lb.
Rostrum and abdomen compressed dorsoventrally; third pair of pereiopods bearing chelae.............................infraorder Astacidea 5
2a(la).
Fingers of chelae of first and second pereiopods with apical tufts of long setae (Fig.lb); supraorbital spine on either side of base of
rostrum; some pereiopods with exopods....................................................................................................................family Atyidae 3
2b.
Fingers of chelae of first and second pereiopods without tufts; no supraorbital spines; all pereiopods lacking exopods ....family Palaemonidae .............................................................................................................................................................................................4
FIGURE 28
(a – d, f) Lateral view of left first pleopods of first-form males (after Hobbs, 1972);
(e) caudal view of same. a, Orconectes i. inermis; b, Procambarus (Remoticambarus) pecki Hobbs;
c, Orconectes propinquus; d, Cambarus latimanus; e, Faxonella creaseri Walls; and f, Procambarus
gracilis.
992
H. H. Hobbs III
FIGURE 29
Dorsal view of chelae and carpus of first form males (after Hobbs, 1972). a, Fallicambarus fodiens; b, Orconectes rusticus; c, Hobbseus cristatus (Hobbs); Distocambarus (Distocambarus)
crockeri Hobbs and Carlson.
3a(2a).
Eyes reduced, without pigment; all pereiopods with exopods; stygobitic ..........................................................................Palaemonias
3b.
Eyes well developed, pigmented; fifth pereiopod without exopod; epigean ..............................................................................Syncaris
4a(2b).
Second pereiopod distinctly longer than first; mandibular palp present; branchiostegal spine on carapace situated distant to anterior
margin; epigean ..........................................................................................................................................................Macrobrachium
4b.
Second pereiopod only slightly longer than first; mandibular palp absent; branchiostegal spine on carapace situated near or on anterior margin; epigean or hypogean ...................................................................................................................................Palaemonetes
5a(lb).
Males lacking hooks on ischia of all pereiopods; first pleopod of male with distal portion rolled to form cylinder; male never
demonstrating cyclic dimorphism; female lacking annulus ventralis ...........................................................................family Astacidae
Pacifastacus
5b.
Males with ischial hooks on one or more of second through fourth pereiopods (Fig. 19); first pleopod of male complexly folded
with sperm tube opening on one terminal element; male always demonstrating cyclic dimorphism; female with annulus ventralis
............................................................................................................................................................................family Cambaridae 6
6a(5b).
Males with hooks on ischia of second and third pereiopods; very small (15 – 33 mm total length) ................subfamily Cambarellinae
Cambarellus
6b.
Males lacking hooks on ischia of second pereiopods; hooks present on ischia of third, third and fourth, or fourth pereiopod
......................................................................................................................................................................subfamily Cambarinae 7
7a(6b).
Mesial surface of flagellum of antennae fringed ...........................................................................................................Barbicambarus
7b.
Mesial surface of flagellum of antennae never fringed ........................................................................................................................8
8a(7b).
Third maxilliped much enlarged; mesial margin of ischium without teeth...................................................................Troglocambarus
8b.
Third maxilliped not conspicuously large; mesial margin of ischium bearing teeth.............................................................................9
9a(8b).
Coxa of fourth pereiopod with caudomesial boss (Fig. 19c) .............................................................................................................10
9b.
Coxa of fourth pereiopod lacking caudomesial boss ........................................................................................................................15
10a(9a).
First pair of pleopods asymmetrical (Fig. 27b) ...................................................................................................Procambarus (in part)
10b.
First pair of pleopods symmetrical (Fig. 27a) ...................................................................................................................................11
11a(10b).
Well-developed hooks on ischia of third and fourth pereiopods, or terminals of pleopods disposed cephalodistally.........................12
11b.
Well-developed hooks on ischia of third pereiopods only, terminals of pleopods never disposed cephalodistally ..............................13
12a(11 a). Shaft of first pleopods straight and terminating in two very short elements directed caudodistally or distally (Fig. 28a) ......................
Orconectes (in part)
23. Decapoda
12b.
993
Shaft of first pleopods straight or bent caudally; if straight, never terminating other than in two very short elements directed caudodistally or distally (Fig. 28b) ..........................................................................................................................Procambarus (in part)
13a(11 b). Opposable margin of dactyl of chela with angular excision in proximal half (Fig. 29a)..................................................Fallicambarus
13b.
Opposable margin of dactyl of chela lacking angular excision in proximal half (Fig 29b) ................................................................14
14a(13b).
First pleopod with terminal elements directed distally (Fig. 28c) ...........................................................................Orconectes (in part)
14b.
First pleopod with terminal elements directed caudally or, rarely, caudodistally (Fig. 28d)...................................................Cambarus
15a(9b).
First pleopod with two terminal elements, one of which at least twice as long as other (Fig. 28e) .........................................Faxonella
15b.
First pleopod with two or more terminal elements, never two with one at least as long as other ......................................................16
16a(15b).
First pleopod terminating in at least three distinct elements (Fig. 28f) ................................................................Procambarus (in part)
16b.
First pleopod terminating in only two distinct elements ...................................................................................................................17
17a(16b).
Dorsal surface of chela studded with crowded small tubercles (Fig. 29c ...............................................................................Hobbseus
17b.
Dorsal surface of chela with tubercles over lateral half widely scattered at most ..............................................................................18
18a(17b).
First pleopod with proximomesial spur (Fig. 27a) ............................................................................................................Bouchardina
18b.
First pleopod lacking proximomesial spur ........................................................................................................................................19
19a(18b).
Carpus of cheliped slender and longer than mesial margin of palm of chela (Fig 29d); rostrum without marginal spines .....................
Distocambarus
19b.
Carpus of cheliped never conspicuously slender and seldom longer than mesial margin of palm (Fig. 29b); if so, rostrum with marginal spines ...........................................................................................................................................................Orconectes (in part)
LITERATURE CITED
Aiken, D. E. 1968. The crayfish Orconectes virilis: survival in a
region with severe winter conditions. Canadian Journal of
Zoology 46:207 – 211.
Aiken, D. E., Waddy, S. L. 1992. The growth process in crayfish. Reviews in Aquatic Sciences 6(3,4):335 – 381.
Albaugh, D. W. 1972. Insectidice tolerances of two crayfish populations (Procambarus acutus) in southcentral Texas. Bulletin of
Environmental Contamination and Toxicology 8:334 – 338.
Alberstadt, P. J., Steele, C. W., Skinner, C. 1995. Cover-seeking
behavior in juvenile and adult crayfish, Orconectes rusticus: effects of darkness and thigmotactic cues. Journal of Crustacean
Biology 15(3):537 – 541.
Ameyaw-Akumfi, C. E. 1979. Appeasement displays in cambarid crayfish (Decapoda, Astacoidea). Crustaceana, Suppl. 5:136 – 141.
Ameyaw-Akumfl, C. E. 1981. Courtship in the crayfish Procambarus
clarkii (Girard) (Decapoda, Astacidea). Crustaceana 40:57 – 64.
Anderson, R. V. 1978. The effects of lead on oxygen uptake in the
crayfish, Orconectes virilis (Hagen). Bulletin of Environmental
Contamination and Toxicology 20:394 – 400.
Andrews, E. A. 1895. Conjugation in an American crayfish. American Naturalist 29:867 – 873.
Andrews, S. C., Dillaman, R. M. 1993. Ultrastructure of the gill epithelia in the crayfish Procambarus clarkii at different stages of
the molt cycle. Journal of Crustacean Biology 13(1):77 – 86.
Atkinson, R. J. A., Taylor, A. C. 1968. Physiological ecology of
burrowing decapods. Symposia of the Zoological Society of
London 59:201 – 226.
Atwood, H. L., Nguyen, P. V. 1995. Neural adaptation in crayfish.
American Zoologist 35:28 – 36.
Avault, J. W., Jr. 1993. A review of world crustacean aquaculture
with special reference to crayfish. Freshwater Crayfish 9:1 – 12.
Baillie, J., Groombridge, B., (Eds.). 1996. 1996 IUCN Red List of
Threatened Animals. The International Union for Conservation
of Nature and Natural Resources, Kelvyn Press, U.S.A.
Barr, T. C. 1968. Cave ecology and the evolution of troglobites, in:
Dobzhansky, T., Hecht, M. K., Steere, W. C., Eds. Evolutionary
Biology 2:35 – 102.
Barr, T. C., Holsinger, J. R. 1985. Speciation in cave faunas. Annual
Review of Ecology and Systematics 16:313 – 337.
Barr, T. C., Kuehne, R. A. 1971. Ecological studies in the Mammoth
Cave system of Kentucky. II. The ecosystem. Annales de Speleologie Revue Trimestrielle, 26:47 – 96.
Bechler, D. L. 1981. Copulatory and maternal – offspring behavior in
the hypogean crayfish, Orconectes inermis inermis Cope and
Orconectes pellucidus (Tellkampf) (Decapoda, Astacidae). Crustaceana 40:136 – 143.
Bechler, D. L. 1983. Contamination of Maramec Spring. North
American Biospeleology Newsletter 29:5 – 6.
Bechler, D. L. 1995. A review and prospectus of sexual and interspecific pheromonal communication in crayfish. Freshwater Crayfish 8:657 – 667.
Beck, J. T. 1980. Life history relationships between the bopyrid isopod Probopyrus pandalicola and one of its freshwater shrimp
hosts Palaemonetes paludosus. American Midland Naturalist
104:135 – 154.
Beck, J. T., Cowell, B. C. 1976. Life history and ecology of the freshwater caridean shrimp, Palaemonetes paludosus (Gibbes).
American Midland Naturalist 96:52 – 65.
Becker, C. D., Genoway, R. G., Merrill, J. A. 1975. Resistance of a
northwestern crayfish, Pacifastacus leniusculus (Dana), to elevated temperatures. Transactions of the American Fisheries Society 104:374 – 387.
Bendell-Young, L., Harvey, H. H. 1991. Metal concentrations in crayfish tissues in relation to lake pH and metal concentrations in water and sediments. Canadian Journal of Zoology 69:1076 – 1082.
Berrill, M., Arsenault, M. 1985. Laboratory induced hybridization of
two crayfish species, Orconectes rusticus and O. propinquus.
Journal of Crustacean Biology 5:346 – 349.
Berrill, M., Hollett, L., Margosian, A., Hudson, J. 1985. Variation in
tolerance to low environment pH by the crayfish Orconectes
994
H. H. Hobbs III
rusticus, 0. propinquus, and Cambarus robustus. Canadian
Journal of Zoology 63:2586 – 2589.
Black, J. B. 1958. Ontogeny of the first and second pleopods of the
male crayfish Orconectes clypeatus (Hay). Tulane Studies in Zoology 6:190 – 203.
Black, J. B. 1975. Inheritance of the blue color mutation in the crawfish Procambarus acutus acutus (Girard). Proceedings of the
Louisiana Academy of Science 38:25 – 27.
Blake, M. A., Hart, P. J. B. 1993. The behavioral responses of juvenile signal crayfish Pacifastacus leniusculus to stimuli from
perch and eels. Freshwater Biology 29(1):89 – 97.
Blank, G. S., Figler, M. H. 1996. Interspecific shelter competition between the sympatric crayfish species Procambarus clarkii (Girard) and Procambarus zonangulus (Hobbs and Hobbs). Journal of Crustacean Biology 16(2):300 – 309.
Borash, J. L., Moore, P. A. 1997. Ultrastructure, morphology and
distribution of sensory structures on the chelae of the crayfish,
Orconectes rusticus, Orconectes virilis and Orconectes propinquus. Ohio Journal of Science 97(2-Program Abstracts):A-11.
Bouchard, R. W., Bouchard, J. W. 1995. Two new species and subgenera
(Cambarus and Orconectes) of crayfishes (Decapoda: Cambaridae)
from the Eastern United States. Notulae Naturae 471:1–21.
Bovbjerg, R. V. 1956. Some factors affecting aggressive behavior in
crayfish. Physiological Zoology 29(2):127 – 136.
Bowman, T. E., Abele, L. G. 1982. Classification of the recent Crustacea, in: Bliss, D. E. Ed., The biology of Crustacea. Vol. 1:Systematics, the fossil record, and biogeography. Academic Press,
New York, pp. 1 – 27.
Boyd, G. L. 1997. Metabolic rates and life history of aquatic organisms inhabiting Logan Cave stream in northwest Arkansas.
Coop Unit Publication No. 29, Arkansas Cooperative Fish and
Wildlife Research Unit, U. S. Geological Survey, Fayetteville,
Arkansas, 107 pp.
Brown, A. V., Pierson, W. K., Brown, K. B. 1994. Organic carbon resources and the payoff-risk relationship in cave ecosystems. Proceedings of the Second International Conference on Ground
Water Ecology, United States Environmental Protection Agency,
Atlanta, Georgia, pp. 67 – 76.
Brown, K. 1981. Low genetic variability and high similarities in the
crayfish genera Cambarus and Procambarus. American Midland Naturalist 105:225 – 232.
Brown, P. B. 1995. Physiological adaptations in the gastrointestinal
tract of crayfish. American Zoologist 35:20 – 27.
Bruski, C. A., Dunham, D. W. 1987. The importance of vision in agonistic communication of the crayfish Orconectes rusticus. I: An
analysis of bout dynamics. Behavior 103:83 – 107.
Budd, T. W., Lewis, J. C., Tracey, M. L. 1979. Filtration feeding in
Orconectes propinquus and Cambarus robustus (Decapoda,
Cambaridae). Crustaceana, Supple. 5:131 – 134.
Burggren, W. W., McMahon, B. R. 1983. An analysis of scaphognathite pumping performance in the crayfish Orconectes virilis:
compensatory changes to acute and chronic hypoxic exposure.
Physiological Zoology 56:309 – 318.
Butler, M. J., Stein, R. A. 1985. An analysis of the mechanisms governing species replacements in crayfish. Oecologia 66:168 – 177.
Caine, E. A. 1975. Feeding and masticatory structures of six species
of the crayfish genus Procambarus (Decapoda, Astacidae).
Forma et Functio 8:49 – 66.
Caine, E. A. 1978. Comparative ecology of epigean and hypogean
crayfish (Crustacea: Cambaridae) from northwestern Florida.
American Midland Naturalist 99:315 – 329.
Capelli, G. M. 1982. Displacement of northern Wisconsin crayfish by
Orconectes rusticus (Girard). Limnology and Oceanography
27:741 – 745.
Capelli, G. M., Hamilton, P. A. 1984. Effects of food and shelter on
aggressive activity in the crayfish Orconectes rusticus (Girard).
Journal of Crustacean Biology 4:252 – 260.
Capelli, G. M., Magnuson, J. J. 1983. Morphoedaphic and biogeographic analyses of crayfish distribution in northern Wisconsin.
Journal of Crustacean Biology 3:548 – 564.
Capelli, G. M., Munjal, B. M. 1982. Aggressive interactions and resource competition in relation to species displacement among
crayfish of the genus Orconectes. Journal of Crustacean Biology
2:486 – 492.
Carpenter, S. R., Kitchell, J. F. 1992. Trophic cascade and biomanipulation: interface of research and management. Limnology and
Oceanography 37:208 – 213.
Carpenter, S. R., Kitchell, J. F. 1993. The trophic cascade in lakes,
Cambridge University Press, Cambridge, United Kingdom.
Cesaroni, D., Allegrucci, G., Sbordoni, V. 1992. A narrow hybrid
zone between two crayfish species from a Mexican cave. Journal of Evolutionary Biology 5:643 – 659.
Chace, F. A., Jr., Hobbs, H. H., Jr. 1969. The freshwater and terrestrial decopod crustaceans of the West Indies with special reference to Dominica. U. S. National Museum Bulletin 292:1 – 258.
Charlebois, P. M., Lamberti, G. A. 1996. Invading crayfish in a
Michigan stream: direct and indirect effects on periphyton and
macroinvertebrates. Journal of the North American Benthological Society 15(4):551 – 563.
Ciruna, K. A., Dunham, D. W., Harvey, H. H. 1995. Detection and
response to food versus conspecific tissue in the crayfish Cambarus bartonii (Fabricius, 1978) (Decapoda, Cambaridae).
Crustaceana 68(6):782 – 788.
Clancy, P. 1997. Feeling the pinch: the troubled plight of America’s
crayfish. Nature Conservancy 47(3):10 – 15.
Claussen, D. L. 1980. Thermal acclimation in the crayfish, Orconectes rusticus and 0. virilis. Comparative Biochemistry and
Physiology 66A:377 – 384.
Cooper, J. E. 1975. Ecological and behavioral studies in Shelta Cave,
Alabama, with emphasis on decapod crustaceans. Ph.D. Thesis,
University of Kentucky, Lexington, 364 pp.
Cooper, J. E., Braswell, A. L. 1995. Observations on North Carolina
crayfishes (Decapoda: Cambaridae). Brimleyana 22:87 – 132.
Cooper, J. E., Cooper, M. R. 1978. Growth, longevity, and reproductive strategies in Shelta Cave crayfishes. National Speleological
Society Bulletin 40:97.
Corey, S. 1987a. Comparative fecundity of four species of crayfish in
southwestern Ontario, Canada (Decapoda, Astacidea). Crustaceana 53:276 – 286.
Corey, S. 1987b. Intraspecific diferences in reproductive potential, realized reproduction and actual production in the crayfish Orconectes propinquus (Girard 1852) in Ontario. American Midland Naturalist 118:424 – 432.
Corey, S. 1990. Life history of Cambarus robustus Girard in the Eramosa-Speed river system of south-western Ontario, Canada
(Decapoda, Astacidae). Crustaceana 59:225 – 230.
Covich, A. P. 1978. Spatial and temporal patterns of foraging activity
among radio-monitored crayfish. Page 13, in: 26th Annual
Meeting of the North American Benthological Society, Titles
and Abstracts.
Covich, A. P. 1982. Crayfish foraging and ecosystem control by selective omnivory. Page 14, in: Payne, J. F., Ed. Crayfish Distribution Patterns. Symposium, American Society of Zoologists,
Louisville, KY.
Covich, A. P., Dye, L. L., Mattice, J. S. 1981. Crayfish predation on
Corbicula under laboratory conditions. American Midland Naturalist 105:181 – 188.
Cox, D. K., Beauchamp, J. J. 1982. Thermal resistance of juvenile
23. Decapoda
crayfish, Cambarus bartoni [sic] (Fabricius): experiment and
model. American Midland Naturalist 108:187 – 193.
Crandall, K. A. 1993. Molecular systematics and evolutionary biology of the crayfish subgenus Procericambarus (Decapoda: Cambaridae). Ph.D. Thesis, Washington University, St. Louis, MO.
Crandall, K. A., Fitzpatrick, J. F., Jr. 1996. Crayfish molecular systematics: using a combination of procedures to estimate phylogeny. Systematic Biology 45(1):1 – 26.
Crawshaw, L. I. 1974. Temperature selection and activity in the crayfish Orconectes immunis. Journal of Comparative Physiology
95:315 – 322.
Creed, R. P., Jr. 1994. Direct and indirect effects of crayfish grazing
in a stream community. Ecology 75(7):2091 – 2103.
Crocker, D. W., Barr, D. W. 1968. Handbook of the crayfishes of Ontario. Miscellaneous Publication of the Royal Ontario Museum.
Uni. of Toronto Press, Toronto, 158 pp.
Crowl, T. A., Covich, A. P. 1990. Predator-induced life-history shifts
in a freshwater snail. Science 247:949 – 951.
Cuadras, J. 1993. Secretory organelles in the crayfish nervous system.
Comparative Biochemistry and Physiology 104A(3):419 – 422.
Daniels, W. H., D’Abramo, L. R., Graves, K. F. 1994. Ovarian development of female Red Swamp Crayfish (Procambarus clarkii) as
influenced by temperature and photoperiod. Journal of Crustacean Biology 14(3):530 – 537.
Daveikis, V. F., Alikhan, M. A. 1996. Comparative body measurements, fecundity, oxygen uptake, and ammonia excretion in
Cambarus robustus (Astacidae, Crustacea) from an acidic and a
neutral site in northeastern Ontario, Canada. Canadian Journal
of Zoology 74:1196 – 1203.
Davies, I. J. 1984. Effects of on experimental whole-lake acidification
on a population of the cray fish Orconectes virilis. North American Benthological Society Abstracts, 32:35.
DeCoursey, P. J. 1983. Biological timing. in: Bliss, D. E., Ed. The biology of Crustacea. Vol. 7, Behavior and Ecology. Academic
Press, New York, pp. 107 – 162.
Deyrup, M., Franz, R. (Eds.). 1994. Rare and Endangered Biota of
Florida. Vol. IV. Invertebrates. Uni. Press of Florida, Gainesville,
798 pp.
Dickson, G. W., Franz, R. 1980. Respiration rates, ATP turnover and
adenylate energy change in excised gills of surface and cave crayfish. Comparative Biochemistry and Physiology 65A:375 – 379.
Dickson, G. W., Briese, L. A., Giesey, J. P., Jr. 1979. Tissue metal concentrations in two crayfish species cohabiting a Tennessee cave
stream. Oecologia 44:8 – 12.
Dickson, G. W., Giesy, J. P. Jr., Briese, L. A. 1982. The effect of
chronic cadmium exposure on phosphoadenylate concentrations and adenylate energy charge of gills and dorsal muscle tissue of crayfish. Environmental Toxicology and Chemistry
1:147 – 156.
Dickson, J. S., Dillaman, R. M., Roer, R. D., Roye, D. B. 1991. Distribution and characterization of ion transporting and respiratory filaments in the gills of Procambarus clarkii. Biological Bulletin 180:154 – 166.
DiDonato, G. T., Lodge, D. M. 1993. Species replacements among
Orconectes crayfishes in Wisconsin lakes: the role of predation
by fish. Canadian Journal of Fisheries and Aquatic Sciences
50:1484 – 1488.
Dimond, J. B., Kadunce, R. E., Gelchell, A. S., Blease, J. A. 1968.
Persistence of DDT in crayfish in a natural environment. Ecology 49:759 – 762.
DiStefano, R. J., Neves, R. J., Helfrich, L. A., Lewis, M. C. 1991. Response of the crayfish Cambarus bartonii bartonii to acid rain
exposure in southern Appalachian streams. Canadian Journal of
Zoology 69:1585 – 1591.
995
Dobkin, S. 1971. The larval development of Palaemonetes cummingi
Chace, 1954 (Decapoda, Palaemonidae), reared in the laboratory. Crustaceana 20:285 – 297.
Doughtie, D. G., Rao, K. R. 1984. Histopathological and ultrastructural changes in the antennal gland, midgut, hepatopancreas,
and gill of grass shrimp following exposure to hexavalent
chromium. Journal of Invertebrate Pathology 43:89 – 108.
Dubé, P., Portelance, B. 1992. Temperature and photoperiod effects
on ovarian maturation and egg laying of the crayfish, Orconectes limosus. Aquaculture 102:161 – 168.
Dunham, D. W., Ciruna, K. A., Harvey, H. H. 1997. Chemosensory
role of antennules in the behavioral integration of feeding by
the crayfish Cambarus bartonii. Journal of Crustacean Biology
17(1):27 – 32.
Dunham, D. W., Oh, J. W. 1992. Chemical sex discrimination in the
crayfish Procambarus clarkii: role of antennules. Journal of
Chemical Ecology 18:2362 – 2372.
Dunham, D. W., Oh, J. W. 1996. Sex discrimination by female Procambarus clarkii (Girard, 1852) (Decapoda, Cambaridae): use
of chemical and visual stimluli. Crustaceana 69(4):534 – 542.
Elliott, W. R. 1997. Conservation of the North American cave and
karst biota. Ecosystems of the World, Subterranean Biota, Elsevier Science, pp. 1 – 24.
Emery, A. R. 1970. Fish and crayfish mortalities due to an internal
seiche in Georgian Bay, Lake Huron. Journal of the Fisheries
Research Board of Canada, 27:1165 – 1168.
Eng, L. L. 1981. Distribution, life history, and status of the California freshwater shrimp, Syncaris pacifica (Holmes). Inland Fish.
Endangered Species Program, Special Publication 81 – 1:1 – 27.
Eversole, A. G. 1995. Distribution of three rare crayfish species in
South Carolina. Freshwater Crayfish 8:113 – 120.
Eversole, A. G., Foltz, J. W. 1993. Habitat relationships of two crayfish species in a mountain stream. Freshwater Crayfish
9:300 – 310.
Feminella, J. W., Resh, V. R. 1989. Submersed macrophytes and
grazing crayfish:an experimental study of herbivory in a Californian freshwater marsh. Holarctic Ecology 12:1 – 8.
Fernández-de-Miguel, F., Aréchiga, H. 1992. Sensory inputs mediating
two opposite behavioural responses to light in the crayfish Procambarus clarkii. Journal of Experimental Biology 164:153 – 169.
Fetzner, J. W., Jr. 1996. Biochemical systematics and evolution of the
crayfish genus Orconectes (Decapoda: Cambaridae). Journal of
Crustacean Biology 16(1):111 – 141.
Fielder, D. D. 1972. Some aspects of the life histories of three closely
related crayfish species, Orconectes obscurus, O. s. sanborni,
and O. propinquus. Ohio Journal of Science 72(3):129 – 145.
Figiel, C. R., Jr., Babb, J. G., Payne, J. F. 1991. Population regulation
in young of the year crayfish, Procambarus clarkii (Girard,
1852) (Decapoda, Cambaridae). Crustaceana 61(3):301 – 307.
Figiel, C. R., Jr., Miller, G. L. 1995. The frequency of chela autotomy
and its influence on the growth and survival of the crayfish Procambarus clarkii (Girard, 1852) (Decapoda, Cambaridae).
Crustaceana 68(4):472 – 483.
Figler, M. H., Twum, M., Finkelstein, J. E., Peeke, H. V. S. 1995.
Maternal aggression in red swamp crayfish (Procambarus
clarkii, Girard): the relation between reproductive status and
outcome of aggressive encounters with male and female conspecifics. Behaviour 132(1,2):107 – 125.
Fitzpatrick, J. F., Jr. 1977. A new crawfish of the genus Hobbseus
from northeast Mississippi, with notes on the origin of the
genus (Decapoda, Cambaridae). Proceedings of the Biological
Society of Washington 90:367 – 374.
Fitzpatrick, J. F., Jr. 1983. A revision of the dwarf crawfishes. Journal
of Crustacean Biology 3:266 – 277.
996
H. H. Hobbs III
Fitzpatrick, J. F., Jr. 1986. The pre-Pliocene Tennessee River and its
bearing on crawfish distribution (Decapoda: Cambaridae).
Brimleyana 12:123 – 146.
Fitzpatrick, J. F., Jr. 1987a. The subgenera of the crawfish genus Orconectes (Decapoda: Cambaridae). Proceedings of the Biological
Society of Washington 100:44 – 74.
Fitzpatrick, J. F., Jr. 1987b. Notes on the so-called “blue color
phase” in North American cambarid crawfishes (Decapoda, Astacoidea). Crustaceana 52(3):316 – 319.
Flint, R. W. 1975. Growth in a population of the crayfish Pacifastacus leniusculus from a subalpine lacustrine environment. Journal of the Fisheries Research Board of Canada 32:2433 – 2440.
Flynn, M. A., Holliday, C. W. 1994. Renal Na, K-ATPase and osmoregulation in the crayfish, Procambarus clarkii. Comparative
Biochemistry and Physiology 107(2):349 – 356.
Flynn, M. F., Hobbs, H. H. III. 1984. Parapatric crayfishes in southern Ohio: evidence of competitive exclusion? Journal of Crustacean Biology 4:382 – 389.
Font, W. F. 1994. Alloglossidium greeri n. sp. (Digenea:Macroderoididae) from the Cajun Dwarf crayfish, Cambarellus schufeldti,
in Louisiana, U.S.A. Transactions of the American Microscopical Society 113(1):86 – 89.
Fotenot, L. W., Platt, S. G., Dwyer, C. M. 1993. Observation on
crayfish predation by water snakes, Nerodia (Reptilia, Colubridae). Brimleyana 19:77 – 82.
France, R. L. 1984. Comparative tolerance to low pH of three life
stages of the crayfish Orconectes virilis. Canadian Journal of
Zoology 62:2360 – 2363.
France, R. L. 1985a. Preliminary investigations of effects of sublethal
acid exposure on maternal behavior in the crayfish Orconectes
virilis. Bulletin of Environmental Contamination and Toxicology 35:641 – 645.
France, R. L. 1985b. Low pH avoidance by crayfish (Orconectes virilis): evidence for sensory conditioning. Canadian Journal Zoology 63:258 – 262.
France, R. L. 1986. Current status of methods of toxicological research on freshwater crayfish. Canadian Technical Report of
Fisheries and Aquatic Sciences No. 1404:1 – 20.
France, R. L. 1992. The North American latitudinal gradient in
species richness and geographical range of freshwater crayfish
and amphipods. The American Naturalist 139(2):342 – 354.
France, R. L., Collins, N. C. 1993. Extirpation of crayfish in a lake
affected by long-range anthropogenic acidification. Conservation Biology 7(1):184 – 188.
France, R. L., Holmes, J., Lynch, A. 1991. Use of size-frequency data
to estimate the age composition of crayfish populations. Canadian Journal of Fisheries and Aquatic Sciences 48(12):
2324 – 2332.
France, R. L., Graham, L. 1985. Increased microsporidian parasitism
of the crayfish Orconectes virilis in an experimentally acidified
lake. Water Air and Soil Pollution 26:129 – 136.
France, R. L., Welbourn, P. M. 1992. Influence of lake pH and
macrograzers on the distribution and abundance of nuisance
metaphytic algae in Ontario, Canada. Canadian Journal of
Fisheries and Aquatic Sciences 49(1):185 – 195.
Franz, R. 1977. Observations on the food, feeding behavior, and parasites of the striped swamp snake, Regina alleni. Herpetologica
33:91 – 94.
Franz, R., Bauer, J., Morris, T. 1994. Review of biologically significant caves and their faunas in Florida and south Georgia. Brimleyana 20:1 – 109.
Franz, R., Lee, D. S. 1982. Distribution and evolution of Florida’s
troglobitic crayfishes. Bulletin of the Florida State Museum, Biological Sciences 28:53 – 78.
Fuller, S. J., Mellen, A. J., Mosher, M. W., Parker, K. E. 1989. Enzymatic comparison of three species of crayfish by cellulose acetate electrophoresis. Transactions Missouri Academy of Science 23:7 – 11.
Gamradt, S. C., Kats, L. B. 1996. Effect of introduced crayfish and
mosquitofish on California newts. Conservation Biology
10(4):1155 – 1162.
Garvey, J. E., Stein, R. A. 1993. Evaluating how chela size influences
the invasion potential of an introduced crayfish (Orconectes
rusticus). American Midland Naturalist 129:172 – 181.
Garvey, J. E., Stein, R. A., Thomas, H. M. 1994. Assessing how fish
predation and interspecific prey competition influence a crayfish
assemblage. Ecology 75(2):532 – 547.
Gelder, S. R. 1996. A review of the taxonomic nomenclature and a
checklist of the species of the Branchiobdellae (Annelida: Clitellata). Proceedings of the Biological Society of Washington
109(4):653 – 663.
Godley, J. S., McDiarmid, R. W., Rojas, N. N. 1984. Estimating prey
size and number in crayfish-eating snakes, genus Regina. Herpetologica, 40:82 – 88.
Gore, J. A., Bryant, R. M., Jr. 1990. Temporal shifts in physical habitat of the crayfish, Orconectes neglectus (Faxon). Hydrobiologia 199:131 – 142.
Gowing, H., Momot, W. T. 1979. Impact of brooktrout (Salvelinus
fontinalis) predation on the crayfish Orconectes virilis in three
Michigan lakes. Journal of the Fisheries Research Board of
Canada 36:1191 – 1196.
Grow, L. 1981. Burrowing behaviour in the crayfish, Cambarus diogenes diogenes Girard. Animal Behaviour 29:351 – 356.
Guiasu, R. C., Barr, D. W., Dunham, D. W. 1996. Distribution and
status of crayfishes of the genera Cambarus and Fallicambarus
(Decapoda: Cambaridae) in Ontario, Canada. Journal of Crustacean Biology 16(2):373 – 383.
Guinot, D. 1994. Decapoda Brachyura. in: Juberthie C., Decu, V.,
Eds. Encyclopedaedia Biospeologica, Tome I, Société de
Biospéologie, Moulis-Bucarest, pp. 165– 179.
Hamr, P., Berrill, M. 1985. The life histories of north temperate
populations of the crayfish Cambarus robustus and Cambarus
bartoni [sic]. Canadian Journal of Zoology 63:2313 – 2322.
Hanson, J. M., Chambers, P. A., Prepas, E. E. 1990. Selective foraging by the crayfish Orconectes virilis and its impact on macroinvertebrates. Freshwater Biology 24:69 – 80.
Hart, C. W., Jr. 1994. A dictionary of non-scientific names of freshwater crayfishes (Astacoidea and Parastacoidea), including
other words and phrases incorporating crayfish names. Smithsonian Contributions to Anthropology No. 38, 127 pp.
Hart, C. W., Jr., Clark, J. 1987. An interdisciplinary bibliography of
freshwater crayfishes (Astacoidea and Parastacoidea) from Aristotle through 1985. Smithsonian Contributions to Zoology No.
455, 437 pp.
Hart, D. G., Hart, C. W., Jr. 1974. The ostracod family Entocytheridae. Academy of Natural Sciences of Philadelphia, Monograph
18, 239 pp.
Hasiotis, S. T. 1995. Notes on the burrow morphologies and nesting
behaviors of adults and juveniles of Procambarus clarkii and
Procambarus acutus acutus (Decapoda: Cambaridae). Freshwater Crayfish VIII:623 – 634.
Hatt, H. 1984. Structural requirements of amino acids and related
compounds for stimulation of receptors in crayfish walking leg.
Journal of Comparative Physiology 155A:219 – 231.
Hayes, W. A., II. 1975. Behavioral components of social interactions
in the crayfish Procambarus gracilis (Bundy) (Decapoda, Cambaridae). Proceedings of the Oklahoma Academy of Science
55:1 – 5.
23. Decapoda
Hayes, W. A., II. 1977. Predator response postures of crayfish. I. The
genus Procambarus (Decapoda, Cambaridae). Southwestern
Naturalist 21:443 – 449.
Hazlett, B. A. 1985. Chemical detection of sex and condition in the
crayfish Orconectes virilis. Journal of Chemical Ecology
2:181 – 189.
Hazlett, B. A. 1990. Source and nature of the disturbance – chemical
system in crayfish. Journal of Chemical Ecology 16(7):
2263 – 2275.
Hazlett, B. A. 1994a. Crayfish feeding responses to zebra mussels depend on microorganisms and learning. Journal of Chemical
Ecology 20(10):2623 – 2630.
Hazlett, B. A. 1994b. Alarm responses in the crayfish Orconectes virilis and Orconectes propinquus. Journal of Chemical Ecology
20(7):1525 – 1535.
Hazlett, B. A., Anderson, F. E., Esman, L. A., Stafford, C., Munro, E.
1992. Interspecific behavioral ecology of the crayfish Orconectes rusticus. Journal of Freshwater Ecology 7:69 – 76.
Hazlett, B. A., Rittschof, D. 1985. Variation in rate of growth in the
crayfish Orconectes virilis. Journal of Crustacean Biology
5:341 – 346.
Hazlett, B. A., Rittschof, D., Rubenstein, D. 1974. Behavioral biology of the crayfish Orconectes virilis. Home range. American
Midland Naturalist 92:301 – 319.
Hedgpeth, J. W. 1949. The North American species of Macrobrachium (River Shrimp). Texas Journal of Science 1(3):28 – 38.
Hedgpeth, J. W. 1968. The atyid shrimp of the genus Syncaris in California. Internationale Revue der Gesamten Hydrobiologie und
Hydrographie 53(4):511 – 524.
Hill, A. M., Lodge, D. M. 1994. Diel changes in resource demand:
competition and predation in species replacement among crayfishes. Ecology 75(7):2118 – 2126.
Hill, A. M., Lodge, D. M. 1995. Multi-trophic-level impact of sublethal interactions between bass and omnivorous crayfish. Journal of the North American Benthological Society 14(2):306 – 314.
Hill, A. M., Sinars, D. M., Lodge, D. M. 1993. Invasion of an occupied niche by the crayfish Orconectes rusticus: potential importance of growth and mortality. Oecologia 94:303 – 306.
Hobbs, H. H., Jr. 1969. On the distribution and phylogeny of the
crayfish genus Cambarus. in: Holt, P. C., Hoffman, R., Hart,
C. W., Jr, Eds. The distributional history of the biota of the
southern Appalachians. Part I: Invertebrates. Research Division
Monograph 1. Virginia Polytechnic Institute, Blacksburg, VA,
pp. 93 – 178.
Hobbs, H. H., Jr. 1972. Crayfishes (Astacidae) of North and Middle
America. Biota of Freshwater Ecosystems. U.S. Environmental
Protection Agency, Water Pollution Control Research Service
Identification Manual 9, 173 pp.
Hobbs, H. H., Jr. 1974. Synopsis of the families and genera of
crayfishes (Crustacea: Decapoda). Smithsonian Contributions to
Zoology No. 164, 32 pp.
Hobbs, H. H., Jr. 1976. Adaptations and convergence in North
American crayfishes. Freshwater Crayfish 2:541 – 551.
Hobbs, H. H., Jr. 1981. The crayfishes of Georgia. Smithsonian Contributions to Zoology No. 318, 549 pp.
Hobbs, H. H., Jr. 1986. Highlights of a half century of crayfishing.
Freshwater Crayfish 6:12 – 23.
Hobbs, H. H., Jr. 1988. Crayfish distribution, adaptive radiation,
and evolution. in: Holdich, D. M., Lowery, R. S., Eds. Freshwater crayfish biology: Management and exploitation. Croom
Helm, London, pp. 52 – 82.
Hobbs, H. H., Jr. 1989. An illustrated checklist of American crayfishes (Decapoda: Astacidae, Cambaridae, and Parastacidae).
Smithsonian Contributions to Zoology No. 480, 236 pp.
997
Hobbs, H. H., Jr., Franz, R. 1986. New troglobitic crayfish with comments on its relationship to epigean and other hypogean crayfishes of Florida. Journal of Crustacean Biology 6:509 – 519.
Hobbs, H. H., Jr., Hall, E. T. Jr. 1974. Crayfishes (Decapoda: Astacidae). in: Hart, C. W., Jr. Fuller, S. L. H., Eds. Pollution ecology
of freshwater invertebrates. Academic Press, New York, pp.
195 – 241.
Hobbs, H. H., Jr., Peters, D. J. 1993. New records of entocytherid
ostracods infesting burrowing and cave-dwelling crayfishes,
with descriptions of two new species. Proceedings of the Biological Society of Washington 106(3):455 – 466.
Hobbs, H. H., Jr., Whiteman, M. 1991. Notes on the burrows, behavior, and color of the crayfish Fallicambarus (F.) devastator (Decapoda: Cambaridae). Southwestern Naturalist 36(1):127 – 135.
Hobbs, H. H., Jr., Hobbs, H. H. III, Daniel, M. A. 1977. A review of
the troglobitic decapod crustaceans of the Americas. Smithsonian Contributions to Zoology No. 244, 183 pp.
Hobbs, H. H., III. 1976. Observations on the cave-dwelling crayfishes of Indiana. Freshwater Crayfish 2:405 – 414.
Hobbs, H. H., III. 1980. Studies of the cave crayfish, Orconectes inermis inermis Cope (Decapoda, Cambaridae). Part IV: Mark-recapture procedures for estimating population size and movements of
individuals. International Journal of Speleology 10:303 – 322.
Hobbs, H. H., III. 1987. Gasoline pollution in an Indiana cave. in:
Program of the 1987 National Speleological Society Convention, p. 33.
Hobbs, H. H., III. 1992. Caves and Springs, in: Hackney, C. T., Adams,
S. M., Martin, W. H. Eds., Biotic communities of the southeastern
United States, Chapter 3: Wiley, New York pp. 59 – 131.
Hobbs, H. H., III. 1993. Trophic relationships of North American
freshwater crayfishes and shrimps. Contributions in Biology and
Geology, Milwaukee Public Museum. Milwaukee, Wisconsin,
110 pp.
Hobbs, H. H., III. 1994. Biogeography of subterranean decapods in
North and Central America and the Caribbean region (Caridea,
Astacidea, Brachyura). Hydrobiologia 287:95 – 104.
Hobbs, H. H., III. 1998. Decapoda (Caridea, Astacidea, Anomura).
in: Juberthie, C., Decu, V. Eds. Encyclopædia Biospeologica. Société de Biospélogie, Moulis-Bucarest, Pages 891 – 911.
Hobbs, H. H., III. 2000 Subterranean Exosystems: Crustacea, in:
Wilkens, H., Culver, D. C., Humphreys, W. F., Eds., Ecosystems
of the World — Subterranean Biota, Chapter 5: Elsevier Science,
The Netherlands pp. 95 – 107.
Hobbs, H. H., III, Jass, J. P. 1988. The crayfishes and shrimp of
Wisconsin (Decapoda: Palaemonidae, Cambaridae). Special
Publications in Biology and Geology, No. 5. Milwaukee Public
Museum. Milwaukee, WI. 177 p.
Hobbs. H. H., III, Jass, J. P., Huner, J. V. 1989. A review of global crayfish introductions with particular emphasis on two North American species (Decapoda:Cambaridae). Crustaceana 56:299 – 316.
Holdich, D. M., Reeve, I. D. 1988. Functional morphology and
anatomy. in: Holdich, D. M., Lowery, R. R. S., Eds. Freshwater
Crayfish:Biology, Management and Exploitation. Timber Press,
Portland, OR, Pages 11 – 51.
Holsinger, J. R. 1988. Troglobites: the evolution of cave-dwelling organisms. American Scientist 76:146 – 153.
Holt, P. C., Opell, B. D. 1993. A checklist of and illustrated key to
the genera and species of the Central and North American
Cambarincolidae (Clitellata:Branchiobdellida). Proceedings of
the Biological Society of Washington 106(2):251 – 295.
Horns, W. H., Magnuson, J. J. 1981. Crayfish predation on lake
trout eggs in Trout Lake, Wisconsin. Rapports et Proces- Verbaux des Reunions, Conseil International pour Exploration de
la Mer 178:299 – 303.
998
H. H. Hobbs III
Hubschman, J. H. 1967. Effects of copper on the crayfish Orconectes
rusticus (Girard). II. Mode of toxic action. Crustaceana 12:
141 – 150.
Hubschman, J. H., Rose, J. A. 1969. Palaemonetes kadiakensis Rathbun: post embryonic growth in the laboratory (Decapoda,
Palaemonidae). Crustaceana 16:81 – 87.
Huner, J. V. 1977. Observations on the biology of the river shrimp
from a commercial bait fishery near Port Allen, Louisiana. in:
31st Annual Conference of the Southeastern Association of Fish
and Wildlife Agencies, Proceedings, Pages 380 – 386.
Huner, J. V. 1995. Ecological observations of red swamp crayfish, Procambarus clarkii (Girard, 1852), and white river crayfish, Procambarus zonangulus Hobbs and Hobbs 1990, as regards their
cultivation in earthen ponds. Freshwater Crayfish 10:456 – 468.
Huner, J. V., Barr, J. E. 1984. Red Swamp Crawfish:Biology and Exploitation. Louisiana Sea Grant College Program. Louisiana
State University, Baton Rouge, 136 pp.
Huppop, K. 1985. The role of metabolism in the evolution of cave
animals. National Speleological Society Bulletin 47:136 – 146.
Huryn, A. D., Wallace, J. B. 1987. Production and litter processing
by crayfish in an Appalachian mountain stream. Freshwater Biology 18:277 – 286.
Jacobson, R. R. 1996. Cave crayfish Cambarus aculabrum recovery
plan. U. S. Fish and Wildlife Service, Southeast Region, Atlanta,
37 pp.
Jacobson, T. R., Hartfield, P. 1997. Alabama cave shrimp Palaemonias alabamae recovery plan. U. S. Fish and Wildlife Service,
Southeast Region, Atlanta, 46 pp.
James, A. 1979. The value of biological indicators in relation to
other parameters of water quality. in: James, A., Evison, L. Eds.
Biological Indicators of water quality. Wiley, Chichester, U K,
Pages 1 – 16.
Jezerinac, R. F., Stocker, G. W., Tarter, D. C. 1995. The crayfishes
(Decapoda:Cambaridae) of West Virginia. Bulletin of the Ohio
Biological Survey, New Series 10(1):1 – 193.
Johannes, R. E., Webb, K. L. 1970. Release of dissolved organic
compounds by marine and freshwater invertebrates. in: Hood,
D. W. Eds. Symposium on organic matter in natural waters. Institute of Marine Sciences. University of Alaska, Fairbanks,
Pages 257 – 273.
Johnson, S. K. 1977. Crawfish and freshwater shrimp diseases. Texas
A&M University, Agricultural Extension Service, College Station, 19 pp.
Johnston, C. E., Figiel, C. 1997. Microhabitat parameters and lifehistory characteristics of Fallicambarus gordoni Fitzpatrick, a
crayfish associated with pitcher-plant bogs in southern Mississippi. Journal of Crustacean Biology 17(4):687 – 691.
Jordan, F., Babbitt, K. J., McIvor, C. C., Miller, S. J. 1996. Spatial
ecology of the crayfish Procambarus alleni in a Florida wetland
mosaic. Wetlands 16(2):134 – 142.
Kasuya, E., Tsurumaki, S., Kanie, M. 1996. Reversal of sex roles in
the copulatory behavior of the imported crayfish Procambarus
clarkii. Journal of Crustacean Biology 16(3):469 – 471.
Keller, T. A. 1992. The effects of the branchiobdellid annelid Cambarincola fallax on the growth rate and condition of the crayfish Orconectes rusticus. Journal of Freshwater Ecology
7(2):165 – 171.
Kershner, M. W., Lodge, D. M. 1995. Effects of littoral habitat and
fish predation on the distribution of an exotic crayfish, Orconectes rusticus. Journal of the North American Benthological
Society 14(3):414 – 422.
Koppelman, J. B., Figg, D. E. 1995. Genetic estimates of variability
and relatedness for conservation of an Ozark cave crayfish
species complex. Conservation Biology 9(5):1288 – 1294.
Kulkarni, G. K., Glade, L., Fingerman, M. 1991. Oogenesis and effects of neuroendocrine tissues on in vitro systhesis of protein
by the ovary of the Red Swamp Crayfish Procambarus clarkii
(Girard). Journal of Crustacean Biology 11(4):513 – 522.
Kushlan, J. A., Kushlan, M. S. 1980. Population fluctuations of the
prawn, Palaemonetes paludosus, in the Everglades. American
Midland Naturalist 103:401 – 403.
Lachaise, F., Le Roux, A., Hubert, M., Lafont, R. 1993. The molting
gland of crustaceans: localization, activity, and endocrine control (a review). Journal of Crustacean Biology 13(2):198 – 234.
Lamontagne, S., Rasmussen, J. B. 1993. Estimating crayfish density
in lakes using quadrats: maximizing precision and efficiency.
Canadian Journal of Fisheries and Aquatic Sciences 50:
623 – 626.
Layne, J. R., Manis, M. L., Claussen, D. L. 1985. Seasonal variation
in the time course of thermal acclimation in the crayfish Orconectes rusticus. Freshwater Invertebrate Biology 4:98 – 104.
Leitheuser, A. T., Holsinger, J. R., Olson, R., Pace, N. R., Whitman,
R. L., Whitmore, T. 1985. Ecological analysis of the Kentucky
cave shrimp, Palaemonias ganteri Hay, at Mammoth Cave National Park (Phase V). Old Dominion University Research Foundation, Norfolk, 102 pp.
Levenbach, S., Hazlett, B. A. 1996. Habitat displacement and the
mechanical and display functions of chelae in crayfish. Journal
of Freshwater Ecology 11(4):485 – 492.
Lichtwardt, R. W. 1986. The Trichomycetes: Fungal Associates of
arthropods. Springer, New York, 343 pp.
Light, T., Erman, D. C., Myric, C., Clarke, J. C. 1995. Decline of the
Shasta crayfish (Pacifastacus fortis Faxon) of northeastern California. Conservation Biology 9(6):1567 – 1577.
Lisowski, E. A. 1983. Distribution, habitat, and behavior of the Kentucky cave shrimp, Palaemonias ganteri Hay. Journal of Crustacean Biology 3:88 – 92.
Lodge, D. M. 1991. Herbivory on freshwater macrophytes. Aquatic
Botany 41:195 – 224.
Lodge, D. M. 1993a. Species invasions and deletions: community effects and responses to climate and habitat change. in: Kareiva,
P. M., Kingsolver, J. G., Huey, R. B. Eds. Biotic interactions and
global change. Sinauer Associates, In., Sunderland, MA, Pages
367 – 387.
Lodge, D. M. 1993b. Biological invasions: lessons for ecology.
Trends in Ecology and Evolution 8:133 – 137.
Lodge, D. M., Brown, K. M., Klosiewski, S. P., Stein, R. A., Covich,
A. P., Leathers, B. K., Bronmark, C. 1987. Distribution of freshwater snails: spatial scale and the relative importance of physicochemical and biotic factors. American Malacological Bulletin
5:73 – 84.
Lodge, D. M., Hill, A. M. 1994. Factors governing species composition, population size, and productivity of cool-water crayfishes.
Nordic Journal of Freshwater Research 69:111 – 136.
Lodge, D. M., Kershner, M. W., Aloi, J. E., Covich, A. P. 1994. Effects of an omnivorous crayfish (Orconectes rusticus) on a
freshwater littoral food web. Ecology 75:1265 – 1281.
Lodge, D. M., Kratz, T. K., Capelli, G. M. 1986. Long-term dynamics of three crayfish species in Trout Lake, Wisconsin. Canadian
Journal of Fisheries and Aquatic Sciences 43:993 – 998.
Lodge, D. M., Lorman, J. G. 1987. Reductions in submersed macrophyte biomass and species richness by the crayfish Orconectes
rusticus. Canadian Journal of Fisheries and Aquatic Sciences
44:591 – 597.
Lodge, D. M., Stein, R. A., Brown, K. M., Covich, A. P., Brönmark,
C., Garvey, J. E., Klosiewski, S. P. 1998. Predicting impact of
freshwater exotic species on native biodiversity: Challenges in
spatial scaling. Australian Journal of Ecology 23:53 – 67.
23. Decapoda
Loring, M. W., Hill, L. G. 1978. Temperature selection and shelter
utilization of the crayfish Orconectes causeyi. Southwestern
Naturalist 21:219 – 226.
Lorman, J. G. 1980. Ecology of the crayfish Orconectes rusticus in
northern Wisconsin. Ph.D. Thesis, University of Wisconsin,
Madison, WI, 227 pp.
Lorman, J. G., Magnuson, J. J. 1978. The role of crayfishes in
aquatic ecosystems. Fisheries, 3:8 – 10.
Lyle, C. 1938. The crawfishes of Mississippi, with special reference to
the biology and control of destructive species. Iowa State College Journal of Science 13:75 – 77.
MacIsaac, H. J. 1994. Size-selective predation on zebra mussels (Dreissena polymorpha) by crayfish (Orconectes propinquus). Journal
of the North American Benthological Society 13(2):206 – 216.
Magnuson, J. J., Beckel, A. L. 1985. Exotic species:A case of biological pollution. Wisconsin Academy Review 32:8 – 10.
Maranhão, P. J. C. Marques, Madeira, V. 1995. Copper concentrations in soft tissues of the red swamp crayfish Procambarus
clarkii (Girard, 1852), after exposure to a range of dissolved
copper concentrations. Freshwater Crayfish 10:282 – 286.
Martin, S. M. 1997. Crayfishes (Crustacea: Decapoda) of Maine.
Northeastern Naturalist 4(3):165 – 188.
Mason, J. C. 1970a. Copulatory behavior of the crayfish, Pacifastacus trowbridgii (Stimpson). Canadian Journal of Zoology
48:969 – 976.
Mason, J. C. 1970b. Maternal – offspring behavior of the crayfish,
Pacifastacus trowbridgi (Stimpson). American Midland Naturalist 84:463 – 473.
Mason, J. C. 1974. Crayfish production in a small woodland stream.
Department of Environmental Fisheries and Marine Sciences,
Pacific Biological Station, Nanaimo, British Columbia. 30 pp.
Mather, M. E., Stein, R. A. 1993. Using growth / mortality trade-offs to
explore a crayfish species replacement in stream riffles and pools.
Canadian Journal of Fisheries and Aquatic Sciences 50:88 – 96.
Mathur, D., Schutsky, R. M., Purdy, E. J. Jr. 1982. Temperature preference and avoidance responses of the crayfish, Orconectes obscurus, and associated statistical problems. Canadian Journal of
Fisheries and Aquatic Sciences 39:548 – 553.
Maude, S. H., Williams, D. D. 1983. Behavior of crayfish in water
currents: hydrodynamics of eight species with reference to their
distribution patterns in southern Ontario. Canadian Journal of
Fisheries and Aquatic Sciences 40:68 – 77.
McGregor, S. W., O’Neil, P. E., Rheams, K. F., Moser, P. H., Blackwood, R. 1997. Biological, geological, and hydrological investigations in Bobcat, Matthews, and Shelta caves and other selected caves in north Alabama. Geological Survey of Alabama
Bulletin 166. 198pp.
McGriff, D. 1983a. The commercial fishery for Pacifastacus leniusculus, from the Sacramento-San Joaquin Delta. Freshwater Crayfish 5:403 – 417.
McGriff, D. 1983b. Growth, maturity, and fecundity of the crayfish
Pacifastacus leniusculus from the Sacromento – San Joaquin
Delta. California Fish and Game 69:227 – 242.
McMahon, B. R., Hankinson, J. J. 1993. Respiratory adaptations in
burrowing crayfish. Freshwater Crayfish 9:174 – 182.
McMahon, B. R., Stuart, S. A. 1995. Simulating the crayfish burrow
environment: air exposure and recovery in Procambarus clarkii.
Freshwater Crayfish 8:451 – 461.
McWhinnie, M. A., O’Connor, J. D. 1967. Metabolism and low temperature acclimation in the temperate crayfish, Orconectes virilis. Comparative Biochemistry and Physiology 20:131 – 145.
Mellon, D., Jr., Sandeman, C., Sandeman, R. E. 1992. Characterization of oscillatory olfactory interneurons in the protocerebrum
of the crayfish. Journal of Experimental Biology 167:15 – 38.
999
Mitchell, D. J., Smock, L. A. 1991. Distribution, life history and production of crayfish in the James River, Virginia. American Midland Naturalist 126(2):353 – 363.
Momot, W. T. 1984. Crayfish production: a reflection of community
energetics. Journal of Crustacean Biology 4:35 – 54.
Momot, W. T. 1992. Further range extensions of the crayfish Orconectes rusticus in the Lake Superior basin of northwestern
Ontario. Canadian Field-Naturalist 106(3):397 – 399.
Momot, W. T. 1995. Redefining the role of crayfish in aquatic ecosystems. Reviews in Fisheries Science 3:33 – 63.
Momot, W. T., Gowing, H. 1977a. Response of the crayfish Orconectes virilis to exploitation. Journal of the Fisheries Research
Board of Canada 34:1212 – 1219.
Momot, W. T., Gowing, H. 1977b. Production and population dynamics of the crayfish Orconectes virilis in three Michigan
lakes. Journal of the Fisheries Research Board of Canada
34:2041 – 2055.
Momot, W. T., Gowing, H., Jones, P. D. 1978. The dynamics of crayfish and their role in ecosystems. American Midland Naturalist
99:10 – 35.
Momot, W. T., Hauta, P. L. 1995. Effects of growth and mortality
phenology on the cohort P/B of the crayfish, Orconectes virilis.
Freshwater Crayfish 8:265 – 275.
Moshiri, G. A., Goldman, C. R., Godshalk, G. L., Mull, D. R. 1970.
The effects of variations in oxygen tension on certain aspects of
respiratory metabolism in Pacifastacus leniusculus (Dana)
(Crustacea:Decapoda). Physiological Zoology 43:23 – 29.
Mundahl, N. D., Benton, M. J. 1990. Aspects of the thermal ecology
of the rusty crayfish Orconectes rusticus (Girard). Oecologia
82:210 – 216.
Naqvi, S. M., Howell, R. D. 1993a. Cadmium and lead uptake by
red swamp crayfish (Procambarus clarkii) of Louisiana. Bulletin
of Environmental Contamination and Toxicology 51:296 – 302.
Naqvi, S. M., Howell, R. D. 1993b. Toxicity of cadmium and lead to
juvenile red swamp crayfish, Procambarus clarkii, and effects
on fecundity of adults. Bulletin of Environmental Contamination and Toxicology 51:303 – 308.
Naqvi, S. M., Leung, T. 1983. Trifluralin and Oryzalin herbicide toxicities to juvenile crawfish (Procambarus clarkii) and mosquitofish (Gambusia affinis). Bulletin of Environmental Contamination and Toxicology 31:304 – 308.
Nelson, D. H., Hooper, D. K. 1982. Thermal tolerance and preference of the freshwater shrimp Palaemonetes kadiakensis. Journal Thermal Biology 7:183 – 187.
Newsom, J. E., Davis, K. B. 1991. Ionic responses of white river
crayfish, Procambarus zonangulus, and red swamp crayfish,
Procambarus clarkii, to changes in temperature and salinity.
American Zoologist 31:43A.
Nevo, E. 1978. Genetic variation in natural populations; pattern and
theory. Theoretical Population Biology 13:121 – 177.
Nielsen, L. A., Reynolds, J. B. 1977. Population characteristics of a
freshwater shrimp, Palaemonetes kadiakensis Rathbun. Transactions of the Missouri Academy Science 10 / 11:44 – 57.
Noblitt, S. B., Payne, J. F. 1995. A comparative study of selected
chemical aspects of the eggs of the crayfish Procambarus clarkii
(Girard, 1852) and P. zonangulus Hobbs & Hobbs, 1990 (Decapoda, Cambaridae). Crustaceana 68(6):695 – 704.
Noblitt, S. B., Payne, J. F., Delong, M. 1995. A comparative study of
selected physical aspects of the eggs of the crayfish Procambarus
clarkii (Girard, 1852) and P. zonangulus Hobbs & Hobbs,
1990 (Decapoda, Cambaridae). Crustaceana 68(5):575 – 582.
Norrocky, M. J. 1984. Burrowing crayfish trap. Ohio Journal of
Science 84:65 – 66.
Norrocky, M. J. 1991. Observations on the ecology, reproduction
1000
H. H. Hobbs III
and growth of the burrowing crayfish Fallicambarus (Creaserinus) fodiens (Decapoda:Cambaridae) in north-central Ohio.
American Midland Naturalist 125(1):75 – 86.
Nystrom, P., Strand, J. A. 1996. Grazing by a native and an exotic
crayfish on aquatic macrophytes. Freshwater Biology
36(3):673 – 682.
Oh, J. W., Dunham, D. W. 1991. Chemical detection of conspecifics
in the crayfish Procambarus clarkii: role of antennules. Journal
of Chemical Ecology 17:161 – 166.
Olsen, T. M., Lodge, D. M., Capelli, G. M., Houlihan, R. J. 1991.
Mechanisms of impact of an introduced crayfish (Orconectes rusticus) on littoral congeners, snails, and macrophytes. Canadian
Journal of Fisheries and Aquatic Sciences 48(10):1853 – 1861.
Page, L. M. 1985. The crayfishes and shrimps (Decapoda) of Illinois.
Illinois Natutal History Survey Bulletin 33:335 – 448.
Page, L. M., Mottesi, G. B. 1995. The distribution and status of the
Indiana crayfish, Orconectes indianensis, with comments on the
crayfishes of Indiana. Proceedings of the Indiana Academy of
Science 104:103 – 111.
Peck, S. K. 1985. Effects of aggressive interactions on temperature selection by the crayfish, Orconectes virilis. American Midland
Naturalist 114:159 – 167.
Peckarsky, B. L., Fraissinet, P. R., Penton, M. A., Conklin, D. J. Jr.
1990. Freshwater macroinvertebrates of northeastern North
America. Cornell Uni. Press, Ithaca, NY, 422 pp.
Pel, X., Moss, F. 1996. Characterization of low-dimensional dynamics in
the crayfish caudal photoreceptor. Nature 379(6566): 618–621.
Penn, G. H., Jr. 1943. A study of the life history of the Louisiana redcrawfish, Cambarus clarkii Girard. Ecology 24:1 – 18.
Perry, W. L., Lodge, D. M., Lamberti, G. A. 1997. Impact of crayfish
predation on exotic zebra mussels and native invertebrates in a
lake-outlet stream. Canadian Journal of Fisheries and Aquatic
Sciences 54(1):120 – 125.
Pflieger, W. L. 1996. The crayfishes of Missouri. Missouri Department of Conservation, Jefferson City, 152 pp.
Pollard, T. G., Larimer, J. L. 1977. Circadian rhythmicity of heart
rate in the crayfish, Procambarus clarkii. Comparative Biochemistry and Physiology 57A:221 – 226.
Power, M. E. 1992. Top-down and bottom-up forces in food webs:
do plants have primacy? Ecology 73:733 – 746.
Powers, L. W. 1977. A catalogue and bibliography to the crabs
(Brachyura) of the Gulf of Mexico. Contribution to Marine Science, 20(Suppl.):1 – 190.
Price, J. O. 1981. Observations on the trophic ecology of the crayfish
Orconectes neglectus chaenodactylus (Decapoda:Astacidae).
ASB Bulletin 28:91 – 92.
Price, J. O., Payne, J. F. 1984. Postembryonic to adult growth and
development in the crayfish Orconectes neglectus chaenodactylus Williams, 1952 (Decapoda, Astacidea). Crustaceana,
46:176 – 194.
Rabeni, C. F. 1985. Resource partitioning by stream dwelling crayfish: The influence of body size. American Midland Naturalist
113:20 – 29.
Rabeni, C. F. 1992. Trophic linkage between stream centrarchids and
their crayfish prey. Canadian Journal of Fisheries and Aquatic
Sciences 49:1714 – 1721.
Rabeni, C. F., Gossett, M., McClendon, D. D. 1995. Contribution of
crayfish to benthic invertebrate production and trophic ecology
of an Ozark stream. Freshwater Crayfish 10:163 – 173.
Reddell, J. R. 1994. The cave fauna of Texas with special reference
to the western Edwards Plateau. in: Elliott, W. R., Veni, G. Eds.
The caves and karst of Texas. A guide-book for the 1994 convention of the National Speleological Society with emphasis on
the south-western Edwards Plateau. National Speleological
Society, Brackettville, Texas, Pages 31 – 48.
Reiber, C. L. 1995. Physiological adaptations of crayfish to the
hypoxic environment. American Zoologist 35:1 – 11.
Reiber, C. L. 1997. Oxygen sensitivity in the crayfish Procambarus
clarkii: peripheral O2 receptors and their effect on cardiorespiratory functions. Journal of Crustacean Biology 17(2):
197 – 206.
Richter, B. D., Braun, D. P., Mendelson, M. A., Master, L. L. 1997.
Threats to imperiled freshwater fauna. Conservation Biology
11(5):1081 – 1093.
Rittschof, D. 1980. Chemical attraction of hermit crabs and other attendants to simulated gastropod predation sites. Journal of
Chemical Ecology 6:103 – 118.
Roell, M. J., Orth, D. J. 1992. Production of three crayfish populations in the New River of West Virginia, USA. Hydrobiologia
228:185 – 194.
Rogers, R., Huner, J. V. 1985. Comparison of burrows and burrowing behavior of five species of cambarid crawfish (Crustacea,
Decapoda) from the Southern University campus, Baton Rouge,
Louisiana. Proceedings of the Louisiana Academy of Science
48:23 – 29.
Roldman, B. M., Shivers, R. R. 1987. The uptake and storage of iron
and lead in cells of the crayfish (Orconectes propinquus) hepatopancreas and antennae gland. Comparative Biochemistry
and Physiology 86C:201 – 214.
Roush, W. 1997. Hybrids consummate species invasion. Science
227:316.
Rutherford, P. L., Dunham, D. W., Allison, V. 1995. Winning agonistic encounters by male crayfish Orconectes rusticus (Girard)
(Decapoda, Cambaridae): chela size matters but chela symmetry
does not. Crustaceana 68(4):526 – 529.
Rutherford, P. L., Dunham, D. W., Allison, V. 1996. Antennule use
and agonistic success in the crayfish Orconectes rusticus (Girard, 1852) (Decapoda, Cambaridae). Crustaceana 69(1):
117 – 122.
Saffran, K. A., Barton, D. R. 1993. Trophic ecology of Orconectes
propinquus (Girard) in Georgian Bay (Ontario, Canada). Freshwater Crayfish 9:350 – 358.
Sandeman, D., Sandeman, R., Derby, C., Schmidt, M. 1992. Morphology of the brain of crayfish, crabs, and spiny lobsters: a
common nomenclature for homologous structures. Biological
Bulletin 183:304 – 326.
Sarver, R. G., Flynn, M. A., Holliday, C. W. 1994. Renal Na, KATPase and osmoregulation in the crayfish, Procambarus clarkii.
Comparative Biochemistry and Physiology 107A(2):349 – 356.
Savino, J. F., Miller, J. E. 1991. Crayfish (Orconectes virilis) feeding
on young lake trout (Salvelinus namaycush): effect of rock size.
Journal of Freshwater Ecology 6(2):161 – 170.
Siewert, H. F., Buck, J. P. 1991. Effects of low pH on survival of crayfish
(Orconectes virilis). Journal of Freshwater Ecology 6(1):87–91.
Smalley, A. E. 1961. A new cave shrimp from southeastern United
States (Decapoda, Atyidae). Crustaceana 3:127 – 130.
Smith, D. G. 1988. Keys to the freshwater macroinvertebrates of
Massachusetts. No. 3: Crustacea Malacostraca (Crayfish,
isopods, amphipods). Publ. No. 15, 236 – 61 – 250 – 2 – 88-CR.
Division of Water Pollution Control, Westboro, MA, 58 pp.
Smith, T. I, Sandifer, P. A., Smith, M. H. 1978. Population structure
of malaysian prawns, Macrobrachium rosenbergii (deMan),
reared in earthen ponds in South Carolina, 1974 – 1976. Proceedings of world Mariculture Society 9:21 – 38.
Stechey, D. P. M., Somers, K. M. 1995. Potential, realized, and actual
fecundity in the crayfish Orconectes immunis from southwestern Ontario. Canadian Journal of Zoology 73:672 – 677.
Stein, R. A. 1975. Sexual dimorphism in crayfish chelae: functional
significance linked to reproductive activities. Canadian Journal
of Zoology 54:220 – 227.
23. Decapoda
Stein, R. A. 1977. Selective predation, optimal foraging, and the
predator – prey interaction between fish and crayfish. Ecology
58:1237 – 1253.
Stein, R. A. 1979. Behavioral response of prey to fish predators. in:
Stroud, R. H., Clepper, H. Eds. Predator – prey systems in fisheries management. Sport Fisheries Institute, Washington, DC
343 – 353 pp.
Stein, R. A., Magnuson, J. J. 1976. Behavior response of crayfish to a
fish predator. Ecology 57:751 – 761.
Stoffel, L. A., Hubschman, J. H. 1974. Limb loss and the molt cycle
in the freshwater shrimp, Palaemonetes kadiakensis. Biological
Bulletin (Woods Hole, Mass.) 147:203 – 212.
Streever, W. J. 1996. Energy economy hypothesis and the troglobitic
crayfish Procambarus erythrops in Sim’s Sink Cave, Florida.
American Midland Naturalist 135(2):357 – 366.
Strenth, N. E. 1976. A review of the systematics and zoogeography
of the freshwater species of Palaemonetes Heller of North
America (Crustacea:Decapoda). Smithsonian Contributions to
Zoology No. 228: 27 pp.
Strenth, N. E. 1994. A new species of Palaemonetes (Crustacea: Decapoda:Palaemonidae) from northeastern Mexico. Proceedings
of the Biological Society of Washington, 107(2):291 – 295.
Strong, D. R. 1992. Are trophic cascades all wet? Differentiation and
donor-control in speciose ecosystems. Ecology 73:747 – 754.
Stuecheli, K. 1991. Trapping bias in sampling crayfish with baited
funnel traps. North American Journal of Fisheries Management
11:236 – 239.
Swerup, C., Rydzvist, B. 1992. The abdominal stretch receptor organ
of the crayfish. Comparative Biochemistry and Physiology
103A(3):423 – 431.
Taketomi, Y., Nishikawa, S., Koga, S. 1996. Testis and androgenic
gland during development of external sexual characteristics of
the crayfish Procambarus clarkii. Journal of Crustacean Biology
16(1):24 – 34.
Taylor, C. A. 1997. Taxonomic status of members of the subgenus Erebicambarus, Genus Cambarus (Decapoda:Cambaridae) east of the
Mississippi River. Journal of Crustacean Biology 17(2): 352 – 360.
Taylor, C. A., Redmer, M. 1996. Dispersal of the crayfish Orconectes
rusticus in Illinois, with notes on species displacement and habitat preference. Journal of Crustacean Biology 16(3):547 – 551.
Taylor, C. A., Warren, M. L. Jr., Fitzpatrick, J. F. Jr., Hobbs, H. H.
III, Jezerinac, R. F., Pflieger, W. L., Robison, H. W. 1996. Conservation status of crayfishes of the United States and Canada.
Fisheries 21(4):25 – 38.
Taylor, R. C. 1983. Drought-induced changes in crayfish populations
along a stream continuum. American Midland Naturalist
110:286 – 298.
Taylor, R. C. 1984. Thermal preference and temporal distribution in
three crayfish species. Comparative Biochemistry and Physiology 77A:513 – 517.
Taylor, R. C. 1985. Absence of Form I to Form II alternation in male
Procamkerus spiculifer (Cambaridae). American Midland Naturalist 114:145 – 151.
Taylor, R. M., Watson, G. D., Alikhan, M. A. 1995. Comparative
sublethal acute toxicity of copper to the freshwater crayfish
Cambarus robustus Astacidae, Decapoda, Crustacea from an
acidic metal contaminated lake and circumneutral uncontaminated stream. Water Research 29(2):401 – 408.
Thacker, R. W., Hazlett, B. A., Esman, L. A., Stafford, C. P., Keller,
T. 1993. Color morphs of the crayfish Orconectes virilis. American Midland Naturalist 129:182 – 199.
Thorp, J. H., Gloss, S. P. 1986. Field and laboratory tests on acute
toxicity of cadmium to freshwater crayfish. Bulletin of Environmental Contamination and Toxicology 37:355 – 361.
1001
Tierney, A. J., Atema, J. 1986. Effects of acidification on the behavioral responses of crayfishes (Orconectes virilis and Procambarus acutus) to chemical stimuli. Aquatic Toxicology 9:1 – 11.
Tierney, A. J., Atema, J. 1988. Behavioral responses of crayfish
(Orconectes virilis and (Orconectes rusticus) to chemical feeding stimulants. Journal of Chemical Ecology 14:123 – 133.
Tierney, A. J., Dunham, D. W. 1984. Morphology of aesthetasc
sensilla in surface and cave dwelling crayfishes. American Zoologist 24:67A.
Tierney, A. J., Thompson, C, S., Dunham, D. W. 1984. Site of
pheromone reception in the crayfish Orconectes propinquus (Decapoda, Cambaridae). Journal of Crustacean Biology 4:554 – 559.
Truesdale, F. M., Mermilliod, W. J. 1979. The river shrimp Macrobrachium ohione (Smith)(Decapoda, Palaemonidae): its abundance, reproduction, and growth in the Atchafalaya River basin
of Louisiana, USA. Crustaceana 36:61 – 73.
Uiska, E., Dunham, D. W., Harvey, H. H. 1994. Cumulative pattern
in pH change alters response to food in the crayfish Cambarus
bartoni [sic]. Canadian Journal of Zoology 72:187 – 190.
Vermeer, K. 1972. The crayfish, Orconectes virilis, as an indicator
of mercury contamination. The Canadian Field-Naturalist
86:123 – 125.
Vernberg, F. J. 1983. Respiratory adaptations. in: Bliss, D. E., Ed.
The biology of crustacea. Vol. 8. Environmental adaptations.
Academic Press, New York, 1 – 42 pp.
Weber, L. M., Lodge, D. M. 1990. Periphytic food and predatory
crayfish: relative roles in determining snail distribution. Oecologia 82:33 – 39.
Weingartner, D. L. 1977. Production and trophic ecology of two
crayfish species cohabiting an Indiana cave. Ph.D. Thesis,
Michigan State University, East Lansing, 323 pp.
Wheatly, M. G. 1995. An overview of electrolyte regulation in the
freshwater crayfish throughout the molting cycle. Freshwater
Crayfish 8:420 – 436.
Wheatly, M. G., de Souza, S. C. R., Hart, M. K. 1996. Related
changes in hemolymph acid – base status, electrolytes, and
ecdysone in intermolt crayfish (Procambarus clarkii) at 23°C
during extracellular acidosis induced by exposure to air, hyperoxia, or acid. Journal of Crustacean Biology 16(2):267 – 277.
Wiens, A. W., Armitage, K. B. 1961. The oxygen consumption of the
crayfish Orconectes immunis and Orconectes nais in response
to temperature and to oxygen saturation. Physiological Zoology
34:39 – 54.
Wiles, P. R., Guan, R. Z. 1993. Studies on a new method for permanently tagging crayfish with microchip implants. Freshwater
Crayfish 9:419 – 425.
Willman, E. J., Hill, A. M., Lodge, D. M. 1994. Response of three
crayfish congeners (Orconectes spp.) to odors of fish carrion and
live predatory fish. American Midland Naturalist 132(1):44 – 51.
Wood, C. M., Rogano, M. 1986. Physiological responses to acid
stress in crayfish (Orconectes): haemolymph ions, acid – base
status, and exchanges with the environment. Canadian Journal
of Fisheries and Aquatic Sciences 43:1017 – 1026.
Word, B. H., Jr., Hobbs, H. H. Jr. 1958. Observations on the testis of
the crayfish Cambarus montanus acuminatus Faxon. Transactions of the American Microscopical Society 77:435 – 450.
Wright, D. A., Welbourn, P. M. 1993. Effects of mercury exposure
on ionic regulation in the crayfish Orconectes propinquus. Environmental Pollution 82:139 – 142.
Wu, S., Brown, S. E. S. 1980. The occurrence of a freshwater shrimp
Palaemonetes paludosus (Gibbes, 1850) (Crustacea:Palaemonidae) in a warm spring of Wellsville, Colorado. Natural
History Inventory of Colorado 5:1 – 10.
This Page Intentionally Left Blank
Glossary
acclimation process of adjustment of a rate function (i.e., respiratory rate) to a change in ambient environmental conditions.
acetabular plates sclerites associated with the genital field in
adults (provisional genital field in deutonymphs) of certain mite taxa bearing the genital acetabula.
acetabulum (acetabula) knob-like structure with a porous cap
associated with the genital field in deutonymph and
adult mites, thought to function as an osmoregulatory
chloride epithelium. Acetabula are strictly or roughly
paired, and usually are borne on acetabular plates, but
may lie in the gonopore or scattered in the integument
flanking the gonopore.
acidosis decline in the pH of the blood through accumulation
of carbon monoxide or anaerobic endproducts.
acrosome a cytoplasmic inclusion in the cell body of a sperm
cell.
Note: Glossary terms were defined by individual authors in reference
to their chapter. Consequently, terms may actually apply to other
taxa and have broader definitions than indicated here.
acute sharp angle at the end.
adductor muscles in bivalve molluscs, the muscles extending
between shell valves, closing the valves on contraction.
adductor muscle scars raised areas or scars left on the interior
of the ostracode shell by the adductor muscles, generally
leave a distinctive pattern of scars.
adenal produced in the parenchyma (rhabdoids).
adenodactyl auxiliary glandular bulb in the male system of
some turbellarian taxa.
adhesive strip thin layer of chitin between the duplicature and
the outer lamella of the ostracode valve.
adnate closely adherent, joined together.
adoral zone of membranelles orderly arrangement of three or
more compound ciliary organelles (membranelles) that
are serially arranged along the left side of the oral area
of a ciliate.
afferent branchial vessel in Mollusca, the blood vessels bringing deoxygenated blood from the visceral mass, foot,
and mantle to the distal ends of the gill filaments.
ala(e) a coarse or fine, wing-like spine, as in the posteriorly directed alae on the lateral surface of the ostracode carapace.
1003
1004
Glossary
allelochemic biochemical substance produced by one species
and released into the environment that subsequently affects the behavior or physiology of another species.
allochthonous substances (or sometimes species) originating
outside the immediate habitats (e.g., carbon produced by
plants located upstream or in the riparian zone); opposite of autochthonous.
allopatric referring to multiple species whose habitat distributions do not overlap.
allorecruitive type of growth of colonial rotifers whereby
young recruited into the colony come predominantly
from other colonies; results in higher genetic diversity of
the colony. Originally defined as Type I colony formation; see autorecruitive.
allozymes enzyme having a specific biochemical function, but
which differs slightly in its primary structure and in its
electrophoretic mobility, from other enzymes coded by
different alleles of this gene
amictic phase in the life cycle of monogonont rotifers in
which reproduction is parthenogenetic only; see mictic
and mixis.
amoeboid amoeba-like, have no fixed shape, and creeping on
surfaces.
amphidelphic with two opposite ovaries that normally join at
the common vagina.
amphids paired sensory organs located on opposite lateral
surfaces of nematodes; variable in position (lip or neck
region) and shape (pore, slit, circular, spiral or cubshaped).
amphimixis union of egg and sperm through ordinary sexual
reproduction, with the gametes derived from separate individuals.
amphoteric type of female rotifers of the class Monogononta
that can produce two types of eggs in the same clutch,
one developing into female (diploid) and the other into
male (haploid) offspring.
anastomosing interconnecting.
ancestrula in ectoproct bryozoans, the original, single zooid
that emerges from a statoblast, differing from subsequent zooids in the colony by its smaller number of tentacles and inability to form statoblasts.
anchoral processes posteriorly directed apodemes of the
gnathosoma (capitulum) in deutonymphs and adult
mites.
angulate having an angle.
anhydrobiosis ability of bdelloid rotifers to undergo desiccation and then be revived at some later time; a cryptobiotic (latent) state in tardigrades induced by loss of water
through evaporation.
annual occurring once per year.
annulate ringed; surrounded by a ring of a different color;
formed into ring-like segments.
annulations transverse striations (trenches) in the cuticle of
nematodes.
annulus ring; in ectoproct bryozoans, the ring of enlarged
chambers, usually gas-filled, on the periphery of floatoblast valves.
annulus ventralis seminal (spermatophore) receptacle of female crayfish.
anoxia lack of dissolved oxygen.
anoxybiosis cryptobiotic (latent) state in tardigrades induced
by low oxygen tensions.
antenna whiplike, paired, generally elongate sensory appendages arising from anterior area of the head (or
cephalothorax); 1-2 pairs of antennae are present in
arthropods.
antennal gland one of pair of complex excretory glands in
many decapods with duct opening on antenna; green
gland.
antennule paired appendage of first cephalic somite of crayfish; sensory, often bearing aesthetascs.
aperture opening to an anterior cavity, as in the opening of a
snail shell from which foot and body protrude.
apex that part of any structure opposite the base by which it
is attached.
apical pertaining to the apex.
apodeme internal extension of thickened cuticle or exoskeleton in some arthropods, often functioning as muscle attachment site.
apomixis form of parthenogenesis in which meiosis is suppressed and offspring are genetically identical to their
mother.
apophyses see pharyngeal or buccal apophyses.
aposymbiotic describing an organism that would normally
have symbionts but the symbionts are not present.
apotypical (apomorphic) derived, or modified, state of a biologic attribute or morphologic character.
apterous without wings.
areoles segment of hairworm cuticle separated from adjacent
segments (areoles) by longitudinal and transverse furrows; may bear granules, bristles, and contain pores.
articulation a joint.
ascus glandular pocket in some turbellarians (Typhloplanidae).
atrium area where two or more passages meet, as in the space
in flatworms where male and female systems open.
attenuate tapering rapidly or abruptly to a point.
aufwuchs German term whose broader North American usage refers to all small, attached (except macrophytes)
and free-moving organisms forming a living film on
aquatic substrates such as rocks, snags, and plants; included are some microinvertebrates, fungi, algae, and
bacteria.
autochthonous substances (or sometimes species) originating
within the immediate habitat, for example, carbon produced by resident phytoplankton; opposite from allochthonous.
autogamy self-fertilization phenomenon (not true sexual reproduction) in ciliates involving a single individual; haploid gametic nuclei are formed and two fuse with each
other in a single individual; see cytogamy.
autorecruitive type of growth of colonial rotifers whereby
young produced by a colony tend to remain in that
colony; characterized by low genetic diversity of the
colony; seeallorecruitive and geminative.
autotoky type of reproduction where progeny are produced
by a single parent (i.e., hermaphroditism or parthenogenesis).
Glossary
autotrophy synthesis of organic compounds from carbon
dioxide using light as the energy source (in photosynthesis) or inorganic chemical compounds (in chemoautotrophy); contrasts with heterotrophy.
axe-head glochidium unionacean (Mollusca) glochidium
larva characterized by a distinctly quadrate shell form.
axenic culture a bacteria-free culture containing only one
species of organism.
axoneme longitudinal bundle of microtubular fibers in flagella, cilia, and axopodia.
axopodium characteristic feeding pseudopodium of Actinopoda, distinguished by needlelike shape, axonemes, and
bidirectional streaming of cytoplasm.
baffles dorsal and/or transverse ridges posterior to mucrones
on interior of the buccal cavity in some eutardigrades.
basal at or pertaining to the base or point of attachment to,
or nearest the main body.
basal bulb swollen, set-off, basal portion of the pharynx (esphagus) of a nematode.
basipod (basipodite) second segment of an arthropod appendage.
basis second segment from proximal end of segmented appendage.
beak raised portion of dorsal margin of the bivalve shell; oldest portion of the shell.
benthos animals living on or near the bottom of an aquatic
habitat.
bident having two points.
bifid cleft, or divided into two parts; forked.
bilamellate divided into two lamellae or plates.
bioturbation disturbance of sediments by an animal.
bivoltine having two broods or generations per year.
bolus a ball.
boss expanded protuberance, as on caudomesial surface of
coxa of fourth pereiopod of male crayfishes.
brachypterous with short or abbreviated wings.
bristles variously shaped cuticular projections arising from
cuticle.
bromeliads common name of a family of plants with cupshaped leaves that can hold water at the base (mostly
tropical species, e.g. the pineapple); protozoa, ostracodes,
and some other invertebrates regularly inhabit these specialized aquatic microhabitats.
brood chamber body space in parent where eggs or embryos
develop; space between the body and carapace of a
cladoceran, where eggs develop.
brosse distinctive “brush” of cilia arising from specialized
short kineties which is present on the anterodorsal surface
of nondividing individuals of certain gymnostome ciliates.
buccal apparatus anterior part of tardigrade foregut, consisting of buccal tube, muscular pharynx, and pair of piercing stylets.
buccal apophyses cuticular thickenings at junction of buccal
tube for insertion of stylet protractor muscles.
buccal ciliature compound ciliary organelles, the bases of
which are associated with the oral area of some ciliates.
buccal cirri in heterotardigrades, paired sensory appendages,
usually short and hair-like, on either side of each
cephalic papilla on the cephalic plate.
1005
buccal tube support median ventral lamina, also called a “reinforcement rod,” from the buccal ring to the midregion
of the buccal tube in some eutardigrades.
bulbous see pharynx.
bursa folds of cuticle on the male nematode tail that assist in
attaching males to females during mating.
bursa (copulatrix) copulatory bursa; receptacle in some female reproductive systems into which donor spermatozoa are deposited.
byssus proteinaceous threads produced from a byssal gland at
the base of the foot, used to attach juvenile or adult molluscan bivalves to hard surfaces.
calcium phosphate concretions small extracellular spherules
found in gills of unionacean mussels made of calcium
phosphate deposited in consecutive lamellae on a proteinaceous matrix.
calotte head region of a Nematomorpha, usually lighter in
color than the adjoining body.
capitulum sclerotized base of a mite’s gnathosoma.
carapace expanded, hard covering of a major body region
(usually head and/or thorax of some arthropods).
cardate type of trophi (jaw structure) found in rotifers which
functions by creating a sucking action.
cardinal teeth massive conical projections formed at the center of the shell-hinge plate which interdigitate to form
the fulcrum on which the valves open and close.
carina sharp, spiral edge or keel on a snail’s shell, usually
along the center of the whorl.
carpus fifth segment from proximal end of segmented appendage.
cauda posterior extension of the idiosoma in adult males of
certain mite taxa, functioning as an copulatory adaptation during spermatophore transfer.
caudal toward the posterior of an organism.
caudal glands glands found in the tail region, as in those of
nematodes that secrete mucus.
caudal rami bifurcated terminal extension of the abdomen in
some arthropods.
cellulase enzyme that degrades cellulose of plant cell walls.
cement glands glands that secrete material to assist fixation of
egg capsules to a substrate, often near female gonopore
in turbellarians.
centroplast central structure in some heliozoa from which axonemes radiate; synonym: central granule, axoplast; a
type of microtubule organizing center.
cephalic pertaining to, or toward, the head.
cephalic papillae large papillae in head region of nematodes;
also, paired sensory appendages on cephalic plate of heterotardigrades, located between internal and external cirri.
cercus (cerci) paired appendage of the last abdominal segment. In dragonflies, one of two pairs of lateral, terminal, triangular sclerites.
cerebropleural ganglion mass of nerve cell bodies near the
mouth.
chaetae see setae.
chaetotaxy number and arrangement of setae on the body
and appendages.
chela claw or pincer; in crayfish, two opposed distal
podomeres (dactyl and propodus) of certain pereiopods.
1006
Glossary
chelate presence of pincer-like, exoskeletal or cuticular structures at the end of an appendage. In mites, a condition of
the pedipalp in deutonymphs and adults of certain taxa
in which tibia bear a dorsodistal seta, often associated
with a projecting tubercle that opposes the dorsal surface
of the tarsus to form a grasping organ.
chelicera in mites, a two-segmented, paired feeding appendage located dorsomedially on the gnathosoma; usually two-segmented, but one-segmented in deutonymphs
and adults of Hydrachnidae; the distal segment, or
“claw,” functions in piercing and tearing the integument
of hosts or prey.
chitin a long chain, polymeric glucosamine containing 6% nitrogen; used in exoskeletons of many arthropods and often complexed with protein or infiltrated with calcium
for strength.
chloride epithelia oval patches (as in insects) that differ from
normal epithelia and are used for osmoregulation.
chrysolaminarin liquid carbohydrate storage form, -1:3linked glucan; characteristic of flagellates in the order
Chrysomonadida; usually in one or more large refractile
vesicles at the posterior of the cell, optically bluish under
strong light; synonym is leucosin.
cingulum one of two ciliated rings present in the typical rotiferan corona (apical end); seetrochus.
circadian rhythm diurnal or daily activity rhythms.
cirrus A in heterotardigrades, paired filamentous sensory appendage at the anterior lateral margin of the scapular
plate, situated slightly dorsal to the clava.
cirrus eversible, spiny male duct; may contain ejaculatory,
prostatic, or accessory ducts and be enclosed in a bulb or
special propulsory sac.
cladogram classification, usually in the form of a branching
diagram, of a group of organisms based on cladistic
principles (i.e., tracing the evolution of shared, derived
characters from ancestral to derived forms).
clasping apparatus in ostracodes, the heavily sclerotized, apparently moveable sclerite articulating with midportion
of the peniferum in a socket near the ventral cardo and
loop of the spermatic tube of entocytherid hemipenes; includes vertical and horizontal rami.
clava in heterotardigrades, a paired sensory appendage,
usually short and broad, at the anterior lateral margin of
the scapular plate, situated slightly ventral to lateral
cirrus A.
claw raptorial structures; in all mite instars, true claws are
paired, curved structures located terminally on tarsi of
the legs.
clear-water phase period, usually in spring, when lake water
becomes unusually clear due to grazing of herbivorous
zooplankton.
clutch group of eggs.
cocoon structure formed by some species to enclose many egg
capsules or embryos.
cohort group of equal-aged individuals whose survivorship
and fecundity are to be followed throughout the lifespan
of the entire group.
columellar strong central support around which a snail shell
coils.
commensal animal or plant that lives on, in, or with another,
sharing its resources but neither parasitic on it nor injured by it.
concentric in Mollusca, describing the growth lines of a snail
operculum that lie entirely within each other (not forming a spiral).
conglutinate mass of unionacean (Mollusca) glochidia larval
bound by mucus; often resembling prey items of
unionacean glochidial fish hosts (increasing chances of
host contact).
conic shaped like a cone.
conjugation in ciliates, a reciprocal fertilization type of sexual
phenomenon occuring only between members of differing mating types and involving temporary or total fusion
of the pair.
contractile vacuole liquid-filled organelle which functions as
an osmoregulator in the cytoplasm of many protozoa.
copepodid larval stage of copepods that follows the first stage
(the nauplius).
copulatory bulb swollen muscular and connective tissue region containing the male copulatory organ and often
also the seminal vesicle and/or prostate.
copulatory organ terminus of male reproductive systems
which delivers spermatozoa into the body of a mate, i.e.
an intromittent organ. When not armed with hard structures, yet capable of protrusion, it is simply called a penis (or penis papilla). Copulatory organs may be
equipped with hard (sclerotized) structures such as a
stylet or cirrus spines.
corneous slightly hardened but still pliable (cornified or
“horny”) proteinaceous material, as in the operculum of
a snail or the modified terminal elements of first
pleopods of first-form male crayfish.
corona apical region of rotifers; usually a ciliated band
around the anterior end, characteristically composed of
two ciliated rings called the trochus and cingulum; in
some forms, ciliation is absent and may be replaced by
long setae which surround the rim of a funnel-shaped
structure called the infundibulum.
cosmopolitanism idea that the same species are found on different continents.
costa rib or ridge on a snail shell that lies transverse to (at
right angles to) the coiling of the whorl.
coxa proximal (closest to body) segment of a segmented appendage.
coxal plate in mites, a sclerite representing the modified basal
leg segment, attached to the venter of the idiosoma in
all instars. Coxal plates may be expanded and fused
with one another to cover large areas of idiosomal integument.
coxoglandularia series of paired glandularia associated with
the coxal plates of mites.
coxopod proximal segment of an arthropod appendage.
creeping welt slightly raised, often darkened structure on
dipteran larvae.
crenulate evenly rounded and often rather deeply curved;
having an irregularly wavy or serrate outline; possessing
a scalloped margin or rounded tooth.
crescent (postanal crescent) crescent-shaped fold between the
Glossary
cloaca and terminus in male hairworms; usually ornamented with setae or bristles.
cross fibers fine criss-cross fibers in the upper cuticular layers
in mermithid nematodes.
cryobiosis cryptobiotic (latent) state in tardigrades induced
by low temperatures.
cryptobiosis latent state in which metabolism comes to a reversible halt; also known as “anabiosis” or “abiosis.”
crystalline style mucopolysaccharide rod containing digestive
enzymes projecting dorsally from the style sac in the
floor of the bivalve stomach where it is secreted; the
crystalline style is rotated by cilia lining the style sac
against the gastric shield on the dorsal side of the stomach.
ctenidium molluscan gill, in bivalves consisting of two demibranchs formed from the reflection of right and left gill
filaments upon themselves.
cupule cup-shaped segment at the base of the club on some
antennae.
cuticular bar external cuticular thickening on the leg, near or
between the claws in some eutardigrades.
cyclomorphosis regular (often seasonal or annual) phenotypic
change in body size, spine length, etc., found in successive generations of zooplankton within a single habitat
(does not apply to morphological changes shown by a
single animal).
cyrtos basket-like structure supporting the cytopharynx of
Ciliophora in the protistan subphylum Cyrtophora.
cytogamy occurrence of autogamy in each member of a pair
of temporarily fused ciliates.
cytolosomes digestive organelles of protozoa, analogous with
food vacuoles, but containing the organelles of the cell,
which are being broken down under conditions of starvation; synonymous with autophagic vacuole or autophagic vesicle.
cytopharynx cell pharynx, leading from the cytostome to site
of food vacuole formation in protozoa.
cytoproct cell anus; permanent site for phagocytosis in some
protozoa.
cytostome cell mouth; permanent site for phagocytosis in
some protozoa.
dactyl most distal segment of the endopod of segmented appendages; smaller, mesially situated, movable finger of
chela.
demibranch portion of the bivalve ctenidium (gill) formed
from a set of filaments reflecting upon themselves; in
each ctenidium, the inner lateral filaments form the inner
demibranch lying next to the visceral mass and the outer
lateral filaments form the outer demibranch lying next to
the mantle.
denticles small, delicate, spinelike projections, usually of shell
material in numerous taxa; also the small teeth-like
processes lining the stoma of some nematodes; small
teeth.
depressed spire a spire that is not raised above the body
whorl of a snail shell.
determinate growth describing an organism that ceases to
grow in size after reaching a certain size or stage; contrasts with indeterminate growth.
1007
detritus decaying organic material.
deutocerebrum portion of a crustacean’s brain controlling antennules and innervating antennae and portions of the
alimentary tract.
deutonymph second nymphal instar in the generalized mite
life cycle and the only active, free-living nymphal instar
in the water mite life cycle.
dextral snail shells with the aperture on right (when viewed
with the apex pointed away from the observer and with
the aperture fully visible).
diapause period of arrested growth and development; often
used to survive harsh environmental conditions.
diel daily
digestive diverticulum in bivalve molluscs, a digestive organ
surrounding the stomach, area in which the final intracellular phase of digestion takes place.
digitate many finger-like extensions of a mollusc’s mantle.
dioecious individual possessing either a male or a female reproductive system; reproductive systems contained in
separate individuals.
diploid having twice the number of chromosomes normally
occurring in a germ cell, most somatic cells are diploid.
discobolocyst extrusome of some flagellated protozoa, especially Chrysomonadida.
discus-shaped round and flat disk with a thin edge and a
thicker middle.
distal toward the end of a structure and away from the central body opposite of proximal.
diurnal recurring every day and/or relating to the daytime.
diverticulum sac-like structure branching from an internal organ such as the intestine.
doliiformis see pharynx.
dormancy state of minimal metabolic activity when growth
ceases, primarily enabling organisms to survive periods
of adverse conditions.
dorsal furrow strip of soft integument separating dorsal and
ventral shields in highly sclerotized adult mites.
dorsalia small platelets on the dorsum of the idiosoma in deutonymphs and adult mites that function as muscle attachment sites.
dorsal plate large sclerite covering the prodorsal region of the
idiosoma in any mite instar, often expanded to cover
part of the opisthosomal dorsum as well.
dorsal shield single large sclerite or a series of closely fitting
platelets covering the entire dorsum of the idiosoma in
deutonymphs and adults of certain mite taxa.
dorsocentralia series of paired, medially located dorsalia in
mites.
dorsoglandularia series of paired glandularia located on the
dorsum of the idiosoma in mites.
dorsolateralia series of paired, laterally located dorsalia in
mites.
dorsum flattened area adjacent to the hinge line and set off
from the lateral surface of the ostracode carapace.
ductus communis in turbellarians, the terminal (or common)
part of the female canal distal to the joining of vitelline
duct(s) and oviduct(s) in ectolecithal systems.
duplicature narrow band of shell material around outer edge
of the proximal face of the epidermis, composed of the
1008
Glossary
same three layers as the outer lamella; only calcified
layer of the proximal covering, lining, or epidermis separated into the list strip, selvage strip, and flange strip of
the ostracode shell.
ecdysis shedding or molting of an older, smaller exoskeleton
in arthropods.
ecdysone the principal molt-stimulating hormone (in several
forms) of arthropods.
ecophenotypic plasticity variation in a species induced by
nongenetic environmental factors.
ectocyst nonliving outer layer of the body wall of bryozoans;
may be sclerotized or gelatinous.
ectolecithal pertains to systems in which the yolk is not incorporated into the oocyte, i.e., part of the female gonad is
a yolk gland (as in higher Turbellaria).
efferent branchial vessel blood vessel in the bivalve gill axis
carrying oxygenated hemolymph from gill filaments to
the longitudinal vessel of the kidney and eventually to
the heart.
ejaculatory complex series of membranous chambers and associated sclerotized framework located distally in the reproductive tract of male adult mites, functioning as a syringe-like organ for compacting masses of spermatozoa,
assembling spermatophores, and expelling them from the
genital tract through the gonopore.
ejaculatory duct (ductus ejaculatorius) terminal part of the
male duct in Turbellaria, sometimes eversible; carries
spermatozoa and usually prostatic secretions through the
copulatory organ.
ejectisome extrusome of some flagellated protozoa, especially
in Cryptophyta.
electromorph organism possessing a set of enzymes with a
specific migration pattern as determined by electrophoretic techniques.
electrostatic (biotic) filter surface of an animal that attracts
charged particles, because of static-electricity-like
charges on the surface.
elytra hardened, shell-like mesothoracic wings of Coleoptera.
emarginate notched; with an obtuse, rounded, or quadrate
section cut from the margin.
empodium medial unpaired, claw-like structure located terminally between the true claws on the tarsi of legs in larvae of all mite taxa, and in deutonymphs and adults of
Stygothrombidiidae.
encystment latent state in aquatic tardigrades in which resistant cysts are produced under deteriorating environmental conditions.
endemic restricted to a particular geographic region.
endites projections or processes from the inner margin of the
inner branch of a branched appendage.
endocrine relating to the system of glands that produce chemical signals (hormones) in the body.
endocyst in ectoproct bryozoans, the inner living tissues of
the body wall, including epidermis, muscle layers, basement membrane, and peritoneum.
endocytosis process by which minute food particles are engulfed by cells into food vacuoles for the final stages of
digestion, as occurs in terminal tubules of molluscan digestive diverticula.
endogenous initiated from within an individual, such as nervous or hormonal cues controlling diurnal rhythmicity in
the absence of external cues.
endolecithal (entolecithal) pertaining to those systems in
which the yolk is incorporated into the oocyte (as occurs
in more primitive Turbellaria).
endopod mesial, or inner branch of an appendage; mesial ramus of biramous appendage, originating on second segment from base.
endosymbiotic describing an organism which currently has
internal symbionts.
epibranchial cavity in bivalve molluscs, the exhalant mantle
cavity formed dorsal to the ctenidium, which carries water leaving the gill to the exhalant siphon; it also receives
the openings of the gonad, kidney, and anus.
epigean referring to surface (above ground) habitats as opposed to hypogean.
epilimnion portion of a lake above the thermocline and located in the limnetic zone.
epineuston organisms living on or slightly immersed in the
air/water interface; see neuston.
ephippium darkened thickened carapace that protects resting
eggs in cladocera.
estivation general term for a physiologic process whereby animals reduce metabolism and activity levels to survive harsh
conditions, such as pond drying for freshwater bivalves.
eulaterofrontal cilia in bivalve molluscs, stiffened cilia extending laterally from the inhalant side of a gill filament
to form a mesh with eulaterofrontal cilia of adjacent gill
filaments on which particles are filtered.
eupathid specialized, thickened seta of mites located on distal, leg segments in all active instars.
euryoecic pertaining to species tolerant of a wide range of
habitats or environmental conditions.
exchange diffusion transport of ions across epithelia such
that diffusion of one ion species down its concentration
gradient is coupled with transport of a second ion in the
opposite direction.
excrescences teeth on internal margins, denticles on the tips,
and talons on the external margins of the horizontal ramus of the entocytherid ostracode hemipenes.
excretophore large vacuolated cells of the gut epithelium of
turbellarians.
excretory pore plate in mites, a sclerite bearing the excretory
pore in larvae, usually expanded to incorporate the bases
of the setae associated with the pore.
exogenous initiated from outside individuals, such as changes
in light intensity controlling valve-gaping activity of a bivalve.
exopod outer branch of a crustacean appendage; lateral ramus of biramous appendage, originating on the second
segment from the base.
exoskeleton hard external supporting structure, often composed of chitin (with or without calcium carbonate); as
in covering of crustaceans.
exploitative competition form of competition in which the
outcome is determined by differential abilities to harvest
a resource and/or by differences in reproductive rates;
competition in which one organism does not directly
Glossary
inhibit another’s access to a limiting resource by behavioral means; compare with interference competition.
extrapallial fluid in bivalve molluscs, the fluid filling the space
(extrapallial space) between the mantle and the shell.
extrapallial space area between the mantle and shell in molluscs.
extrusome organelle of the pellicle of protozoa, which extrudes a substance or structure, usually in response to a
stimulus.
eye plate in mites, a sclerite bearing one or eyes.
facilitation indirect food web effects which span two trophic
levels; for example, an increase in the abundance of algae because a predator removes a snail herbivore.
facultative an optional process.
fascicles small tooth-like bundle of fibers.
fenestra in ectoproct bryozoans, the clear central portion of a
floatoblast valve.
file row, as in file of cilia or setae.
filiform thread-like; slender and of equal diameter.
filopodium long, filamentous pseudopodium of certain Rhizopoda (amoeba), sometimes branching out without
anastomoses; may function in both food capture and locomotion.
finger guard extension of the ventral cardo along the side of
the dorsal and ventral fingers of the entocytherid ostracode hemipenes.
first-form male one of two morphological forms of cambarine
crayfish; sexually functional male; at least one terminal
element of first pleopod usually corneous.
flagellum multiarticulate, endopodite of antennae in some
arthropods; whiplike locomotory structure of many protozoa.
flange distal ridge of the contact margin of the duplicature
between the list and groove, it may be a part of the outer
lamella of the ostracode shell.
flange groove that part of the duplicature surface of the ostracode valve between the selvage and flange.
flange strip part of the duplicature that forms the flange
groove and the flange of the ostracode valve.
floatoblast in ectoproct bryozoans, a type of statoblast having
a ring of gas-filled chambers; in some cases, the ring becomes buoyant only after being dried.
follicular organ arranged in several small compartments (follicles), i.e., neither diffuse nor compact.
foot a highly muscular locomotory or digging structure.
forcipate type of trophi (jaw structure) found in rotifers having an action like forceps in which the trophi are projected from the mouth to grasp prey.
fossorial associated with digging or burrowing.
fragmentation spherules in Mollusca, the shed tips of digestive cells lining the terminal tubules of digestive diverticulum which are released after intracellular digestion has
been completed; they contain undigested material remaining in food vacuoles and are carried on ciliary rejection tracts in the digestive diverticulum back into the
stomach where their breakdown releases the undigested
contents of their food vacuoles to be egested.
frontal glands glands at anterior terminal region of turbellarians (frontal organ in Acoela).
1009
fulcrate type of aberrant trophi (jaw structure) found in rotifers in the order Seisonidae.
fulcrum one of seven pieces comprising trophi of rotifers; together with the paired rami, they form the incus subunit.
funiculus in ectoproct bryozoans, a tubular strand of tissue
joining the blind end of the gut to the inner colony wall;
the site of statoblast and sperm formation
furca forked posterior end of some gastrotrichs. Also, an appendage-like structure that is the termination of the body
proper in cypridid and cytherid ostracodes; it is articulated to the body and may assist in locomotion but is not
considered a true thoracic leg.
fusiform spindle-shaped; tapering at each end.
gastric mill posterior end of the cardiac stomach of crustaceans and some other arthropods where food is
ground for later digestion.
gastric shield chitinous plate on the dorsal side of the stomach of bivalves (Mollusca) against which the distal tip of
the crystalline style rotates.
geminative type of growth of colonial rotifers whereby young
produced by a colony on any day are all born within a
span of a few hours and leave the parent colony together
as a free-swimming larval colony, attaching to a new
substrate in concert. As a consequence of this phenomenon, the genetic diversity of the colony is low; see autorecruitive and allorecruitive.
gena (genae) the cheek; part of the head on each side below
the eyes, extending ventrally to the gular suture.
genet genetically distinct individual; often used to describe individuals that commonly grow as colonies in which the
individual is not easily distinguished as a separate genetic
entity; contrasts with ramet.
geniculate jointed to permit bending at an abrupt angle; distinctly elbowed.
genital bay area between the posterior coxal groups that accommodates the genital field in adult mites.
genital field area of the idiosoma occupied by the gonopore
and genital acetabula in adult mites.
genital flaps paired movable flaps which close to cover the
gonopore in adults of certain mite taxa; they typically
also cover the genital acetabula, but in some derivative
groups, the acetabula lie on the flaps.
genital papillae papillae situated on the surface of the cuticle
or on elongate finger-like projections on the tail of male
nematodes.
germarium an ovary.
germovitellarium organ containing both oogonia and
oocytes, i.e., ovary and vitelline cells.
gibbosity hump, marked by convexity or swelling, protuberant structure normally found on the dorsal part of an ostracode carapace.
gill axis in Mollusca, portion of the ctenidium from which
gill filaments extend and which is attached to the dorsal
mantle wall.
gill filaments in bivalve molluscs, long, thin, fused lateral extensions from the gill axis which extend ventrally and
then reflect dorsally to form the inner and outer demibranchs.
glandularium (glandularia) specialized organ located in the
1010
Glossary
integument of deutonymphs and adult mites, consisting
of a small, goblet-shaped sac, or “gland,” with a tiny external opening, and an associated seta. Glandularia have
both a sensory and a secretory function.
globose globe-shaped.
glochidium bivalved larval stage of unionacean bivalves
(Mollusca), generally parasitic on fish.
glossa median terminal (or subterminal) lobe of the labium in
some arthropods.
glycocalyx “hairy” secreted layer covering the outer cell
membrane of many of the “naked” rhizopod amoebae.
gnathosoma in mites, anterior region of the body comprising
the mouth and associated appendages, the chelicerae and
pedipalps.
gonochoristic two sexes; males and females in the same population reproducing by cross-fertilization; contrasts with
hermaphroditism.
gonopore external opening of the reproductive system in
adults.
green glands paired excretory glands in crustaceans; also
known as antennal glands.
gubernaculum sclerotized structure that supports and guides
the spicules out of the cloacal opening during mating.
gula throat sclerite, forming the central part of the head beneath the genae.
gullet in flagellated protozoa, anterior invagination from
which the flagella emerge.
gyttja watery, organic sediment that consists mainly of excretory material from benthic animals.
haploid having the number of chromosomes characteristic of
the mature germ cell (half the number of the somatic cell).
haptocyst extrusome of some predatory Ciliophora (Sectoria)
used to capture prey.
heliotropism tendency of certain plants or other organisms to
turn or bend under the influence of light, especially sunlight.
helocrene seepage area where spring water percolates through
a substratum of mosses, leaf litter, and detritus.
hemelytra mesothoracic wings of heteropteran insects.
hemipenes paired penes.
hemocoel internal body cavity formed by the expansion of
the vascular system.
hemocyanin copper-based respiratory pigment which is dissolved within the hemolymph of crustaceans.
hemoglobin iron-containing blood respiratory pigment;
found in a few invertebrates living in aquatic habitats
with low oxygen levels.
hemolymph the circulatory fluid or blood of animals with
open circulatory systems.
hemolymph sinuses or hemocoels open cavities in which the
blood or hemolymph bathes the tissues directly, allowing
exchange of gases, ions, nutrients, and wastes.
hermaphroditic individual containing both male and female
reproductive organs.
heterotrophy consuming, rather than synthesizing, organic
compounds for metabolic activities; contrasts with autotrophy.
hibernaculum thick-walled resting bud occurring in certain
gymnolaemate bryozoans species.
hindgut portion of intestine lying between midgut and rectum, primarily involved with consolidation of undigested
particles into feces.
hinge section of dorsal shell valve in bivalve molluscs containing the teeth and forming the fulcrum on which
valves open and close.
hinge ligament elastic proteinaceous portion of the shell of bivalve molluscs secreted by the mantle isthmus which
connects the mineralized shell valves; it is flexed or compressed when the valves are closed and when adductor
muscles relax and functions to open the valves by returning to an unflexed state.
hinge teeth interdigitating mineralized projections of shell in
bivalve molluscs on the dorsal margins of the valves that
hold the valves in position and serve as the fulcrum on
which they open and close.
histophages protozoa that feed on the tissue of dead metazoa,
as opposed to a parasite feeding on live hosts.
holophyletic group monophyletic clade including all species
descended from a common ancestor.
homoplasy parallel adaptive modification of homologous attributes or structures in species of different clades due to
similar selective pressures.
hooked glochidia glochidia larvae in bivalve molluscs with
spiny hooks on their ventral shell margins.
hormone chemical signal produced and used inside an individual’s body.
hyaline margin clear ectoplasmic material surrounding and/or
preceding the granular body mass of some amoebae during locomotion.
hybrid result of a mating between organisms with different
genotypes; used also for matings between species.
hypersaline term generally referring to a water body whose
salinity is higher than seawater (35 mg / L).
hypodermal cords expansions of the hypodermis between
muscle fields of nematodes.
hypogean subterranean; burrow, cave, or groundwater environment.
hypolimnion portion of a lake below the thermocline and located in the limnetic zone.
hyponeuston organisms living just below the air/water interface of a water body; see neuston.
hyporheic zone groundwater present below and to the side of
a stream and located close enough to be affected by the
movement of stream surface waters (unlike the phreatic
zone).
hypostome pertaining to the ventral portion of the ostracode
head is located a keel-shaped structure which forms the
mouth.
hypoxia condition of environmental oxygen concentration
below air oxygen saturation levels but not necessarily
anoxic.
hysterosoma region of idiosoma of mites posterior to level of
primitive sejugal furrow.
idiosoma body proper of mites, comprising the fused
cephalothorax and abdomen.
imagochrysalis quiescent transitional instar between the deutonymph and adult in the water mite life cycle, representing the suppressed tritonymph.
Glossary
incudate type of trophi (jaw structure) found in rotifers,
which functions by grasping prey with a forceps-like action.
incus subunit of a rotifer trophi composed of fulcrum and
paired rami.
induction change in development or behavior due to receiving
an internal or external signal (such as a hormone or
kairomone)
infauna sediment-dwelling animals; also, but more rarely, animals that burrow into and live inside another organism.
infraciliature total assemblage of all kinetosomes and the associated subpellicular microfibrular and microtubular
structures in ciliate protozoa.
infundibulum corona of rotifers of the order Collothecacea.
inner lamella thin layer of chitin, which, together with the
duplicature, forms a proximal covering of the epidermis
in ostracodes; at inner limit, it is folded back on itself to
form the body wall of the pouch-shaped body of the animal.
instar any developmental stage between molts.
interareolar furrows network of transverse and longitudinal
trenches separating areoles in hairworms; may bear
pores or support bristles and warts.
interference competition form of competition in which one
organism directly or indirectly interferes with another
organism (e.g., by aggression), thus preventing it from
obtaining a limiting resource; forms of interference include physical combat, the threat of combat, noxious
chemical compounds, etc.
interlamellar space cavity formed between ascending and descending gill filaments in Mollusca; water flowing
through the ostia of gill filaments enters the interlamellar
space to be carried dorsally to the epibranchial cavity;
also called the water tube or water channel.
interstitial microhabitat comprising spaces between particles
of submerged sand and gravel in groundwater and hyporheic habitats.
intracytoplasmic lamina filament layer of varying thickness
present within the syncytial integument of rotifers and
acanthocephalans.
ischiopodite third segment from the base of a segmented appendage.
isthmus portion of nematode pharynx between the metacorpus and basal bulb, usually surrounded by the nerve
ring. In bivalve molluscs, the constricted dorsal section
of the mantle separating the mantle into two lateral
shell-secreting halves; the isthmus secretes the proteinaceous hinge ligament connecting the valves.
iteroparous reproducing repeatedly at multiple times during
an organism’s life (cf., semelparous).
kairomone chemical signal produced by a predator that affects behavior, development, or life history of another
species so as to result in higher relative survival of individuals of that species that react to the signal.
karst terrain underlain by fractured and extensively dissolved
carbonate rock (e.g., limestone) and typified by numerous sinkholes, springs, caves, and few surface streams.
kinetid kinetosome and its associated pellicular structures in
Ciliophora, including the cilium, the elementary repeat-
1011
ing structure of the ciliate cortex; the polykinetid is an
organellar complex composed of two or more kinetids.
kinetocyst extrusome of Actinopoda, which secretes materials
that probably serve to make the surface of an axopod
sticky.
kinetoplast DNA-rich organelle near the anterior of some flagellated protozoa (Kinetoplastida), associated with their
unique large mitchondrion.
kinetosome subpellicular structure forming the base of a cilium, but may be without a cilium; it is characterized by
nine triplets of microtubules at right angles to the plasma
membrane arranged in a circle.
kinety single, structurally and functionally integrated (somatic) row of kinetosomes along with their cilia (cilia
not necessarily associated with each inetosome), i.e., a
line of kinetids.
knob in bivalve molluscs, a high rounded major protuberance
with the sides joining the rest of the valve at a distinct
line and at a steep angle.
K-selection selection for life-history characteristics that increase fitness in stable environments where populations
reach densities near environmental carrying capacity; associated with high levels of intraspecific competition.
labial palps pair of thin flattened structures extending from
either side of the mouth region of bivalve molluscs to lie
against the outer surface of the outer demibranch and
the inner surface of the inner demibranch; they function
to receive filtered particles from the ctenidia and sort
them into those accepted for ingestion and those rejected
as pseudofeces.
lacustrine see lentic.
lamellae layers; flattened, leaf-like structures in several phyla,
such as the gills of bivalve molluscs and the buccal platelike projections on the mouth (buccal) ring of some eutardigrades.
Lansing effect putative negative effect of elevated calcium on
survivorship in rotifers in which calcium accumulation in
older mothers is passed on to their offspring; seeorthoclone.
lateral coxal apodeme conspicuous apodeme indicating the
line of fusion between the second and third coxal plates
in larvae of certain mite taxa.
lateral field track usually containing several longitudinal striations running on both sides of the nematode.
lateral teeth elongated lamellar hinge teeth in bivalve molluscs forming anterior and posterior to the cardinal teeth
or posterior to the pseudocardinal teeth in unionacean
bivalves.
LC50 concentration of a toxic agent at which 50% of a cohort will perish within a certain predetermined period;
also called the median lethal concentration and LD50.
lentic standing water environments (e.g., lakes, ponds, wetlands, and both permanent and temporary pools).
limnetic open water, deeper areas of a lake or pond and away
from the shoreline littoral zone.
line of concrescence proximal line of junction of the duplicature and outer lamella of the ostracode valve, the inner
border of the chitin adhesive strip.
lira rib or ridge running along the whorl of a snail shell in the
direction of the coiling.
1012
Glossary
list proximal ridge on the contact margin of the duplicature
of the ostracode valve, not always present.
list strip part of the duplicature from the inner margin to and
including the list of the ostracode valve.
littoral zone or body of water shallow enough for growth of
rooted aquatic vegetation.
lobopodium blunt pseudopodium used in feeding and/or locomotion.
lophophore organized structure of ciliated tentacles used for
capturing suspended food particles from the water, and
probably also providing an important surface for gas exchange; occurring in ectoprocts and certain other phyla.
lorica thickened body wall of rotifers, organized as a syncytial integument possessing two keratin-like proteins
which compose a layer called the intracytoplasmic lamina. A loose-fitting surface covering secreted and/or assembled by protozoa; generally synonymous with test.
lotic running water environments (e.g., creeks, rivers, and
some springs).
lunule crescent or half-moon shaped structure at the base of
each double claw in some eutardigrades.
lyrifissure (lyriform fissure) slender sac-like structures in mites
opening through slits in the integument, apparently functioning as proprioceptors, in all active instars.
lysosome membrane-bound organelle containing hydrolytic
enzymes; in protozoa, they are involved in intracellular
digestion.
macroinvertebrate organisms larger than microinvertebrates
and meiobenthos; retained on coarse sieves with a mesh
2 mm.
macronucleus in ciliates, the nucleus active in transcription
and thus responsible for the phenotype and regulation of
metabolism; see micronucleus.
macrophagous in protozoa, a species that consumes large
items singly by phagocytosis.
macrophyte macroscopic photosynthetic organism growing
submersed, floating, or emergent in water; most are angiosperms but also includes some nonvascular plants and
macroalgae.
macropterous with fully formed wings.
malleate type of trophi (jaw structure) found in certain
monogonont rotifers in which the rami are massive and
may possess teeth along the inner margin.
malleoramate type of trophi (jaw structure) found in monogonont rotifers of the order Flosculariacea, resembling
the malleate form, except in number of teeth on the unci.
malleus subunit of rotifer trophi composed of an uncus and
manubrium.
mandible hardened mouthparts (cephalic appendages) in
many arthropods that are used to grasp, tear, and push
food into the mouth; most anterior gnathal appendage
and just behind the antennae.
mandibular palp endopodite segment of the mandible.
mantle thin extension of the dorsal body wall of molluscs underlying the shell and responsible for its secretion.
mantle cavity space formed between the body and surrounding
mantle tissues in molluscs; completely surrounds the body
in bivalves.
manubrium one of seven pieces comprising the trophi of
rotifers; a manubrium together with an uncus form the
malleus subunit.
marsupium space formed for incubation of embryos and larval stages.
mastax muscular pharynx of rotifers containing jaws called
trophi.
maxillae first or second pair of mouthparts (cephalic appendages) posterior to the mandibles; crustaceans have
two pairs of maxillae, insects have one pair.
maxillary glands paired excretory glands of crustaceans.
maxilliped one of pair of three sets of gnathal appendages situated immediately posterior to second pair of maxillae.
maxillule first maxilla.
medial coxal apodemes series of short apodemes at the medial edges of coxal plates in some larval mites.
medial eye plate (frontal plate) sclerite bearing medial eye in
early derivative mite taxa; the medial eye may be absent
in highly derived groups.
meiobenthos small benthic organisms that pass through a 2
mm mesh but are retained on a 40 – 200 m mesh.
melanin black, brown, or red pigment made of polymerized
amino acids (tyrosine); used for photoprotection and
making structures stronger.
merus fourth segment from proximal end of segmented appendage.
mesial (mesal) pertaining to the middle; toward the middle.
metachronal occurring one after another, in succession.
metacorpus swollen median portion of nematode pharynx,
located between the procarpus and isthmus.
metamere homologous body segment, a somite.
metasome portion of the copepod body anterior to the major
body articulation.
microaerophilic protozoa with aerobic metabolism living in
microhabitats with very low oxygen concentrations.
micronucleus nuclei of ciliates mediating sexual recombination and reproduction, typically much smaller than the
macronucleus; diploid; often multiple.
microphagous protozoa that consume relatively small items
by phagocytosis, collecting many into a single food
vacuole.
microplankton plankton that are 20 – 200 m in size.
microsome simple membrane-bound organelle containing oxidative enzymes; synonymous with microbody.
microvilli microscopic projections from a cell surface that increase the surface area of the cell.
mictic (mixis) phase in the life cycle of monogonont rotifers
in which reproduction is sexual, resulting in a diapausing embryo called a resting egg; seeamictic.
midden refuse heap; commonly, shells of unionacean mussels
that had been consumed by humans or other mammals
and then left at one site.
middle piece structure in sperm cells lying between the head
and flagellum containing mitochondria.
midgut portion of the intestine between stomach and
hindgut; in molluscs, it is characterized by the presence
of the typhlosole, the ciliated surfaces of which sort particles into those returned to the stomach and those
passed to the hindgut for consolidation into feces and
egestion.
Glossary
mixotroph organism driving energy from both autotrophy
and heterotrophy, including those whose autotrophy is
via endosymbionts.
molting process of shedding the cuticle at periodic intervals
to allow expansion of the body in a newly developed
and larger exoskeleton.
monoecious individual possessing both male and female reproductive systems, at least during part of its life cycle.
monophyletic group phylogenetic grouping of species descended from a common ancestor and which are classified together within a single taxonomic unit; compare
with polyphyletic.
monopodial describing an unbranched, cylindrical type of
pseudopodium.
morph variant form within a species.
morphotype morphologically distinct form; in rotifers, a form
present within a single population whose polymorphic
feature(s) (e.g., spines, body size) undergo a seasonal
phenotypic change.
mouth hook vertically-oriented, mandible-like structure in
dipteran larvae.
muciferous body variety of mucocyst found in some Phytomonadida.
mucocyst extrusome of protozoa responsible for secretion of
mucous-like material onto the cell surface.
mucrones small cuticular teeth on interior of the buccal cavity in some eutardigrades.
multispiral growth rings of an operculum of a snail comprised of many, slowly enlarging spirals.
multivoltine having more than two broods or generations per
year.
muscle scars area marking the former position of attachment
of a muscle on the interior of a shell or other support
structure, distinguished from the rest of the shell by a
surrounding groove or by being higher or depressed.
mutualism association between two organisms in which both
benefit.
nacre or nacreous layer inner shell layer of most molluscs, secreted by the mantle as successive layers of calcium carbonate crystals (absent in the Sphaeriidae).
nannoplankton plankton that range from 2 – 20 m in size.
nauplius earliest larval stage of many crustaceans.
neonate newly-born animal.
net growth efficiency percentage of assimilated energy represented by that allocated to tissue growth and gamete
production.
neuston organisms living at or very near the air/water interface of a water body; see hyponeuston, epineuston, and
pleuston.
niche partitioning differences in use of resources that alleviate
interspecific competition.
node on the ostracode carapace, a protuberance smaller than
a lobe or knob but larger than a tubercle, may have the
form of a half sphere.
nonrespired assimilation portion of assimilated energy or
food not catabolized for maintenance energy and,
therefore, available for production of new tissue or gametes.
nymphochrysalis quiescent transitional instar between the
1013
larva and deutonymph in the water mite life cycle, representing the suppressed protonymph.
oblique slanting; neither parallel nor perpendicular to a reference point.
obtuse blunt angle at the end.
ocellus simple eye consisting of a single, bead-like lens, occurring singly or in small groups; distinct from a compound
eye.
omnivore consuming food at more than one trophic level,
e.g., crayfish eat live animals (carnivory) and plants (herbivory) and will also consume dead animals and plants
(detritivory).
open circulatory system circulatory system characterized by
blood not being continually maintained in vessels; rather,
the blood bathes the tissues directly in large hemocoel sinuses before returning to the heart.
operculate formed as a cover.
operculum lid or covering structure, as in some insects.
Horny or calcareous disk on the foot of a snail that covers the aperture when foot is withdrawn; used as protection for predators.
opsiblastic egg type of resting egg produced by gastrotrichs;
see tachyblastic egg.
orifice opening; in ectoprocts, the terminal opening in a zooid
through which the polypide extends.
orthoclone clone of a rotifer species established after many
generations of culturing the ith offspring of every generation (seeLansing effect).
osmobiosis cryptobiotic (latent) state in tardigrades induced
by elevated osmotic pressures.
ostia pores; openings penetrating fused gill filaments in bivalve molluscs with eulamellibranch ctenidia which allow passage of water across the filaments into the interlamellar space.
outer lamella in ostracodes, a hard shell material that covers
the outside of the epidermis that secreted it, composed of
a thick layer of calcite enclosed between two layers of
chitin.
ovate (ovoid) somewhat oval in shape.
oviduct duct from the ovary, through which ova pass.
oviparous females that produce eggs that undergo embryonic
development and hatch outside the body of the female;
see viviparous and ovoviviparous.
ovoviviparous females which produce eggs that undergo embryonic development inside the female and that hatch
just before or soon after being released; see oviparous
and viviparous.
paddles feather-shaped locomotory structures attached anteriorly in Polyarthra rotifers.
pallial pertaining to the pallium or mantle cavity of a mollusc.
pallial line line of muscle scars on the outer perimeter of the
inside of the bivalve mollusc shell, marking points at
which pallial muscles attach the mantle to the shell.
pallial sinus posterior indentation of the pallial line on the inside of the bivalve mollusc shell, marking the space into
which the siphons are withdrawn on valve closure.
palmate like the palm of the hand, with fingerlike processes.
palpal lobes broad, paired, movable lobes on the distal end of
the prementum in odonate insects.
1014
Glossary
palustrine relating to wetland habitats.
papilla one of many small discrete protuberances (as on the
ostracode shell surface) each with steep sides; smaller
than tubercles.
papillate many lobe-like extensions.
paraglossa lateral terminal lobe of the labium.
paramylon reserve food material characteristic of euglenid
flagellates; occurs in a variety of shapes, usually with a
laminated structure.
paraphyletic group monophyletic grouping including only
some of the species descended from a common ancestor.
paratene repeated kinetidal patterns at right angles to the longitudinal axis of a ciliate’s body; superficially gives impression that the ciliary rows run circumferentially
rather than longitudinally in the affected part of the
body.
parietal inside wall of a snail aperture.
paroral membrane synonymous with undulating membrane.
parthenogenesis sexual reproduction in which egg development occurs without fertilization. Amictic parthenogenesis is sexual reproduction without recombination of genetic material.
paucispiral growth lines of a snail operculum, present as a
few, rapidly enlarging spirals.
pectinate having narrow parallel projections suggestive of the
teeth of a comb.
pedal referring to the foot.
pedal feeding use of the foot in bivalve molluscs to feed on
organic detrital deposits in or on sediments; generally involves ciliary tracts on the foot to bring detritus to the
palps or gill food grooves.
pedal gland cement glands in the foot region of rotifers used
to achieve temporary (in pelagic or littoral zooplankton)
or permanent (in sessile forms) attachment to a surface.
pedicles lateral extensions of the corona of certain bdelloid
rotifers.
pedipalp feeding appendage; in mites, the pedipalp typically
has five movable segments, located laterally on the
gnathosoma in all instars. Pedipalps have both sensory
and raptorial functions.
peduncle pseudopodium-like cytoplasmic extension emerging
from the sulcus in some dinoflagellates and used in feeding. Also, general name for a holdfast organelle in diverse groups of free-living protozoa.
pelagic see limnetic.
pellicle surface layer of protozoa, including the plasma membrane and associated membranous, microfibrillar and
microtubular structures and organelles, but excluding
nonliving external coverings such as tests or loricas.
peniferum portion of the entocytherid ostracode copulatory
apparatus that bears the penis (usually its distal portion),
and its proximal portion forms a hinge on which the entire apparatus may be turned through an arc of 180 degrees about the zygum; includes the dorsal and ventral
cardo, costa, spermatic tube, and penis.
penultimate next to last.
pereiopods thoracic appendages of many crustaceans; usually
having a prime locomotory function (walking or swimming) and one or more secondary roles.
perennial living for several years.
periblast part of a statoblast valve of ecotoprocts that excludes the capsule; in floatoblasts, the periblast includes
both annulus and fenestra.
peribuccal lobes flattened cuticular projections surrounding
the mouth of some eutardigrades.
peribuccal (oral) papillae elongated cuticular projections surrounding the mouth of some eutardigrades.
peribuccal papulae short, distinct cuticular projections surrounding the mouth of some eutardigrades.
pericardial cavity remnant of the coelomic cavity surrounding
the heart ventricle in molluscs.
pericardial gland specialized tissue in the wall of the heart auricles of molluscs through which hemolymph fluid is ultrafiltered into the pericardial space from which it enters
the kidney.
pericardium epithelial layer lining the pericardial cavity.
periostracum proteinaceous outer layer of mollusc shells secreted by the mantle edge.
periphyton microalgae growing in a thin layer on mostly vascular plants; sometimes used loosely for microalgae
growing on other substances.
peristalsis rhythmic waves of contraction which moves material through the gut.
peristome area surrounding the cytostome of ciliates, especially where it is specialized relative to the rest of the
body surface.
peristomium first functional segment, usually containing the
mouth, in segmented worms.
phagocytosis process by which a cell engulfs a particle by extending its plasma membrane to form a vacuole or invagination.
phagotrophy nutrition by phagocytosis.
pharyngeal apophyses cuticular thickenings at the junction of
the buccal tube and lumen of the pharynx, alternating in
position with the placoids.
pharyngeal intestinal junction; in nematodes, a cellular valve
located between the pharynx and intestine; may protrude
down into the intestine.
pharynx portion of the alimentary tract located between the
stoma and intestine, often called the esophagus. In turbellarians, “pharynx simplex” is a mostly short, unfolded
tube; “pharynx plicatus” is a protrusible, more or less
tubular fold enclosed in the pharynx cavity; “pharynx
bulbosus” is a bulb separated from the surrounding tissue
by a muscular septum. Pharynx bulbosus is represented
by three types: rosulatus, doliiformis, and variabilis.
Pharynx rosulatus is mostly round with a more or less
vertical axis; pharynx doliiformis is barrel-shaped, anteriorly situated and with a horizontal axis; pharynx variabilis is weakly muscular with variable shape and weakly
differentiated septum.
phasmids paired sensory organs located laterally in the caudal area of nematodes.
phenotypic plasticity expression of different morphologies depending on environmental influences or signals and expressed by a single genotype.
pheromone chemical messenger substance secreted by one individual into the external environment, which elicits a
Glossary
specific response in another individual of the same
species.
photodamage damage caused to molecules due to ultraviolet
radiation in sunlight.
photoperiod length of the light period during a 24 hour lightdark cycle.
photoprotection protection from ultraviolet light, as by a
dark pigment that absorbs light.
phototaxis locomotion directed by light.
phreatic zone flowing groundwater located far enough from
surface streams so that water movement in the streams
does not influence movement of the phreatic waters; seehyporheic zone.
phyllopod a leaf-like or lobed, biramous appendage in some
non-malacostracan crustaceans.
phytophilic aquatic herbivores with a preference for consuming algae; also called algivores.
phytoplanktivorous pertaining to feeding on phytoplankton
(i.e., suspended algae).
phytoplankton unicellular and multicellular algae and
cyanobacteria suspended in the water column.
picoplankton plankton 0.2 – 2 m in size, primarily bacteria.
pinocytosis process by which cells ingest dissolved or colloidal material, involving attraction of these substances
to cell surfaces and then internalizing them through invagination of the plasma membrane.
piptoblast type of statoblast in bryozoans that has no annulus and is not cemented to the colony substrate; formed
only in the family Fredericellidae.
placoids cuticular thickenings in the lumen of the tardigrade
pharynx (pharyngeal bulb); large anterior ones are
macroplacoids, small posterior row are microplacoids.
plankton eukaryotic and prokaryotic organisms living all or
part of their lives in the water column of aquatic ecosystems and at the mercy of water currents.
planospiral having shell whorls confined to a single plane;
coiled but without an elevated spire.
plasma membrane unit or cell which is the outer-most living
layer of protozoa.
plasmodium multinucleate mass of protoplasm enclosed by a
plasma membrane; nonhomologous trophic stage in the
life cycle of some protozoa.
plastron semipermanent bubble of air through which carbon
dioxide from respiration is exchanged with oxygen from
the water.
pleopod one of five pairs of appendages on first five segments
of a decapod’s abdomen; also called “swimmerets,” or
modified into male gonopods.
plesiotypical (plesiomorphic) conservative, or unmodified,
state of a biologic attribute or morphologic character.
podomere leg segment of an arthropod.
polykinetid see kinetid.
polykinety compound ciliary organelle composed of several
kineties, e.g., a cirrus.
polyphyletic group taxonomic assemblage of species that are
not all descended from a common ancestor.
polypide retractable portion of a bryozoan zooid comprising
the central ganglion and all structures related to feeding
and digestion.
1015
polyploid having more than twice the number of chromosomes in somatic cells.
polytrophic see omnivore.
polyvoltine with several generations during the season; contrasts with univoltine.
pore canals in nematomorphs, pores connected to fine tubes
in the cuticle; in ostracodes, a passage through the entire
valve in which are located sensory hairs to the nerves of
the hypodermis.
postcloacal crescent see crescent.
posterior ridge external ridge on unionacean (Mollusca)
shells extending from the umbos posterioventrally to the
shell margin.
postocularia pair of prominent setae located on the dorsum
of the idiosoma posterior to the level of the eyes in deutonymphs and adult mites.
prehensile fitted or adapted for grasping, holding, or seizing.
prehensile palp appendage modified to grasp, hold, or seize;
generally formed of two podomeres, the proximal one
referred to as the prodopus, distal one referred to as the
dactylus of some crustaceans such as ostracodes.
prementum distal segment of the labium in odonate insects,
which bears the palpal lobes.
preocularia pair of prominent setae located on the dorsum of
the idiosoma anterior to the level of the eyes in deutonymphs and adult mites.
preparasitic attendance phoretic association by larvae of
some species of water mites with the preimaginal stage
of their insect host.
prismatic layer molluscan shell layer composed of elongated
calcium carbonate crystals oriented at a 90° angle to the
surface plane of the shell and lying between the outer periostracum and inner nacreous layer.
procorpus anterior portion of the nematode pharynx between
the stoma and metacorpus.
prodelphic with a single anterior-directed ovary in nematodes.
profundal deep zone of a lake below the littoral and open
water photic zone; often restricted to benthic habitats
but may include pelagic zone.
proleg process or appendage that serves the purpose of a leg
but is not a true leg.
promitosis form of nuclear division wherein the endosomal
body in the nucleus divides and forms two polar masses
with a spindle between them; characteristic of some
naked amoeba of the class Heterolobosea.
propodosoma region of a mite’s idiosoma posterior to the
level of the primitive sejugal furrow.
propodus penultimate segment (sixth from base) of a segmented appendage.
proprioceptor sensory receptor that responds to physical or
chemical stimuli originating from the organism.
prostate cells, glands, or organs in male reproductive systems,
releases secretions that mix with sperm.
prostaglandins local chemical mediators inducing short-lasting effects in adjacent tissues.
prostomium region anterior to mouth in some oligochaete
worms.
protandry sequential hermaphroditism with the male stage
preceding the female stage.
1016
Glossary
protocerebrum part of a crustacean’s brain that normally innervates eyes, sinus gland, frontal organs, and head muscles.
protoconch shell of newborn snail or “spat.”
protonephridia excretory-osmoregulatory system of certain
pseudocoelomates, including gastrotrichs and rotifers,
comprised of flame cells and tubules.
protonymph first nymphal instar in the generalized mite life
cycle.
protopodite basal part of a typical crustacean limb consisting
of two, more-or-less consolidated segments and bearing
at its distal extremity an exopodite, endopodite, or both.
proventriculus large cavity in collothecid rotifers that is an
extension of the mastax in which prey are held prior to
being passed into the stomach.
provisional genital field region of the idiosoma occupied by
the rudimentary gonopore and genital acetabula in mite
deutonymphs.
proximal toward the median plane of a body.
pseudobranch “false gill” present in ancylid and planorbid
snails; conical extension of epithelium used in respiration, analogous to ctenidium of prosobranchs.
pseudocardinal teeth in bivalve molluscs, compact shell
lamellae on the anterior portion of the hinge plate of
unionacean mussel shells which performs function of
cardinal teeth.
pseudocoelom body cavity not lined with mesodermal tissue,
as is a true coelom, and found in gastrotrichs, nematodes, and rotifers.
pseudofeces masses of mucous-bound particles rejected from
the labial palps of bivalve molluscs before ingestion and
release into the mantle cavity to be carried out the
siphon on water currents induced by valve clapping (i.e.,
rapid valve adduction).
pseudopodium in protozoa, a dynamic extension of the cytoplasm used in feeding or locomotion; in insects, a soft,
foot-like appendage that is less prominent than a proleg.
pseudosexual eggs in cladocera, asexually produced eggs that
resemble those produced by a sexual process. In rotifers,
diploid resting egg supposedly produced in rotifers by
parthenogenesis in the absence of males; seemixis.
pseudostome aperature of a test or shell of a testate amoeba;
pseudopodia protrude from it during locomotion and/or
feeding.
punctae small pit-like opening of a pore canal on the ostracode shell. Microscopic pores traversing the mineral portion of the shells of sphaeriid bivalve molluscs to open
beneath the periostracum; contain extensions of the
pyramidal cells.
pustule protuberance with a pore in the middle, similar to a
volcano with a crater, about the size of a tubercle on the
ostracode carapace.
pyramidal cells in bivalve molluscs, mantle epithelial cells
that extend through shell punctae (pores) to lie just below the periostracum.
Q10 difference in metabolic rate over a 10°C temperature
range (e.g., Q10 would equal 2 if the respiration rate
doubled between 10 and 20°C).
Q10 (acc.) factor by which a biologic rate function changes
over a 10°C increase in acclimation temperature.
quadrate square or rectangular in shape.
radial pore canal passage through the adhesive strip, between
the duplicature and the outer lamella of the proximal
valve edge of an ostracode.
ramate type of trophi (jaw structure) found in bdelloid rotifers in which the rami are large and semicircular in
shape and unci possess many teeth.
ramus one of the seven pieces comprising rotifer trophi;
paired rami together with the fulcrum form the incus
subunit.
receptaculum seminis part of the female reproductive system
of turbellarians in which recipient spermatozoa are
stored.
reniform suggesting a kidney in shape.
respiratory plate also referred to as branchial plate, modified
from the exopodite and may contain numerous feathered
setae.
resting egg thick-walled cyst enclosing a diapausing, diploid
embryo, usually resistant to extreme environmental conditions. In rotifers these “eggs” are produced by sexual
reproduction.
reticulopodium threadlike, repeatedly branching and anastomosing pseudopodium, functioning more often in food
gathering than locomotion.
retrocerebral organ structure of unknown function in rotifers
composed of the unpaired retrocerebral sac and paired
subcerebral glands together; possible function as an exocrine gland lubricating the anterior part of the body.
retrocerebral sac unpaired glandular structure near the subcerebral gland in the head region of rotifers possessing a
forked duct that opens into the corona.
rhabdions cuticular segments of the nematode stoma, may be
fused and produce an entire stomal wall or separate and
exhibit a segmented wall.
rhabdos organelle supporting the cytopharynx of some ciliates,
especially in the subphylum Rhabdophora; although like a
cyrtos, rhabdos is considered evolutionarily the more
primitive organelle.
rheocrene flowing spring with a substratum of sand and
gravel.
rheotaxis orientation to current (can be positive or negative).
riparian living on the bank of a lake, pond, or stream.
rostrum any extended portion of the head end of an invertebrate.
r-selection selection for life-history traits increasing fitness in
an unstable habitat where catastrophic population reductions prevent high levels of intraspecific competition.
scapular setae second most anterior series of sensory setae on
the dorsal surface of the idiosoma of water mites, consisting of an internal pair (si) and an external pair (se).
sclerite hardened area of the insect body wall bounded by
sutures or membranes.
semivoltine having one brood or generation every two years.
sanguivorous feeding on blood.
scabrous having small, round, raised scales or cones.
second-form male one of two morphological forms of male
cambarine crayfishes; sexually nonfunctional male
lacking corneous terminal elements on first pleopod
(gonopod).
Glossary
sediment focusing uneven deposition of sediments in a basin
so that certain portions of the basin accumulate sediments at a greater rate; usually this occurs in the deepest
parts of the basin.
sejugal furrow primitive line of demarcation on idiosoma at
level between insertions of second and third pairs of legs
in all mite instars, but expressed only in certain early derivative taxa.
selvage middle and principal ridge of the contact margin of
the duplicature of the ostracode valve.
selvage groove part of the duplicature surface between the list
and selvage of ostracode valve.
selvage strip part of the duplicature forming the selvage and
selvage groove of the ostracode valve.
semelparous breeding only once during an organism’s life;
one massive reproductive effort.
seminal receptacle (spermatheca) sac-like structure in the female reproductive system that receives and stores spermatozoa.
seminal vesicle saclike structure in the male reproductive system that stores spermatozoa.
sensillum epithelial sense organ.
septulum cuticular thickening in the lumen of the pharynx,
posterior to the placoids, in the same plane as the pharyngeal apophyses in some eutardigrades.
septum dividing wall or membrane; in ostracodes, a small
ridge on the list strip on the duplicature of the valve.
serial homologies structures having the same embryonic and
evolutionary origin but which differ segmentally (e.g., all
appendages of a crayfish, except the first antennae, arose
from similar embryonic primordia and from a series of
similarly formed appendages in an ancient ancestral
stock).
serotonin chemical messenger stimulating active transport of
sodium ions from the medium in freshwater bivalve molluscs.
serrate notched or toothed on the edge.
sessoblast relatively large statoblast cemented firmly through
the body wall to the substratum; formed only in the bryozoan family Plumatellidae.
setae (or chaetae) epidermal, hair-like extensions of the cuticle.
sexual dimorphism differences in shape and/or size between
females and males of the same species.
sheath in nematodes, a cuticle retained from the last molt
which may surround the entire body or head region.
simplex stage molting stage in tardigrades characterized by
the absence of the buccal apparatus.
sinistral snail shells with the aperture opening toward the left
(when viewed with the apex pointed away from the
observer and with the aperture fully visible).
siphon in bivalve molluscs, tubular structure formed by
fusion of opposite mantle margins directing inhalant (inhalant siphon) and exhalant (exhalant siphon) respiratory and feeding water currents into and out of the mantle cavity. An elongated breathing tube in some insects.
solenidion (solenidia) specialized setiform chemosensory structure in mites; solenidia are located on distal segments of
the pedipalps and legs in all active instars; though seta-like
in appearance, solenidia differ structurally and ontogeneti-
1017
cally from setae and are usually given distinctive designations in chaetotactic formulae.
somatic cell cell that become differentiated into tissues, organs; opposed to embryonic germ cell.
somite homologous body segment, a metamere.
spear protrusible pointed, solid rod-like structure in the
mouth region of some nematodes.
species richness number of species in a habitat.
spermatogenesis formation of sperm.
spermatophore packet of spermatozoa enclosed in a membranous capsule, produced by adult male mites and either
deposited on the substratum or transferred to the female
gonopore.
spermatozoan individual haploid cell produced by males for
sexual reproduction.
sphincter circular muscle forming a ring around a tube, such
as the gut, that closes the tube by contracting.
spicules internal support and/or protective structures of
sponges composed principally of silica or calcium salts.
Also, paired sclerotized structures in the tail of the male
nematode.
spindle snail shell with a thick middle and a tapered point at
both the apex and the aperture ends.
spinneret in nematodes, a terminal pore out of which passes
secretions of the caudal glands.
spiracle respiratory opening connected to the tracheal (respiratory) system of insects.
standing crop biomass amount of tissue produced per unit
area at a set point in time, usually measured as g carbon,
Kcal, or ash free dry mass (AFDM) per m2.
statoblast in phylactolaemate bryozoans, an asexually produced capsule composed of two convex valves enclosing a
mass of yolky material and undifferentiated tissue; capable of germinating a new zooid after obligate dormancy.
statoconia in bivalve molluscs, many small mineral inclusions
in the cavity of a statocyst (gravity orientation organ).
statocyst gravity-orientation organ containing mineral inclusions (statolith or statoconia) in a vesicle lined with sensory cilia.
statolith mineral inclusion in the cavity of a statocyst.
stenopod unbranched, segmented walking leg in some crustaceans, particularly decapods.
stenothermal tolerating little variation in temperature.
stigmata paired external openings of the tracheal system, located between bases of the chelicerae in all active instars
of mites.
stolon in certain ectoproct bryozoans, a cylindrical stem-like
structure from which individual zooids grow at intervals.
stoma portion of the alimentary tract between the oral opening (mouth) and pharynx.
stream order method for classifying streams in which the
smallest permanent streams is designated as a “first order” stream; two first orders streams combine to form a
second order stream; a first and second in combination
are still considered a second order, but when two second
order tributaries join, a third order stream results.
striations (striae) parallel grooves; also, the spiral, microscopic grooves along the whorls of a snail shell (sometimes incorrectly used to refer to spiral ridges or lirae).
1018
Glossary
stygobitic species occur exclusively in hypogean habitat.
stygophilous species occurring in both surface and subterranean waters but with a preference for the latter habitat.
style sac evagination of the stomach floor of molluscs, which
secretes the crystalline style.
stylet sclerotized tube or channel of variable shape and complexity; a hollow, sclerotized, hard, rod-like structure in
the mouth region of some nematodes.
stylet knobs enlarged protuberances (usually three) at the
base of the nematode stylet, usually serve for attachment
of muscles that aid in stylet extension.
stylostome mucopolysaccharide feeding tube secreted by parasitic larval water mites into the body of their host.
subcerebral glands paired glands near the retrocerebral sac in
the head region of rotifers with ducts that lead to the
corona; seeretrocerebral organ.
subitaneous egg one developing and hatching with no arrest
in development.
sulcus depression or furrow on the lateral surface of the organisms in several phyla (e.g., Ostracoda and Mollusca);
in protozoa, a longitudinal groove on the surface of dinoflagellates from which the flagella and, if present, the
peduncle or other feeding structures emerge.
suture region where two structures (e.g., plates or valves) are
joined.
symbiont one member of a symbiotic relationship.
sympatric two or more species whose habitat distributions
overlap.
syngamy fusion of haploid gametes or gametic cells in sexual
reproduction to form a diploid zygote; phenomenon exhibited by some members of all major protozoan taxa
except ciliates.
tachyblastic egg type of egg produced by gastrotrichs in which
development occurs immediately; see opsiblastic egg.
tagma or tagmata body regions (especially in arthropods)
consisting of metameres grouped or fused to perform
similar functions, e.g., head, thorax, and abdomen of a
crustacean or insect.
test loose surface covering of protozoa, created by the inhabitant with secreted and, sometimes, gathered materials;
generally synonymous with lorica, but test is normally
applied to Rhizopoda.
thermocline zone in a lake of relatively rapid temperature
change over a small range of depth; commonly defined
as 1°C change per meter of depth.
tocopherol (a-tocopherol or vitamin E) organic compound
that has been shown to induce mixis (sexual reproduction) in some rotifers and lengthen the lifespan in others.
torpidity condition or quality of being dormant or inactive, a
temporary loss of all or part of the power of sensation or
motion.
transverse muscle attachment scar conspicuous muscle attachment scar in the posterior region of the third coxal
plates in larvae of certain mite taxa.
trichocyst type of extrusome emitting a spindle-shaped, nontoxic projectile.
triploid having three times the haploid number of chromosomes.
tritocerebrum portion of a crustacean’s brain that principally
innervates the antennae and a section of the alimentary
tract.
tritonymph third nymphal instar in the generalized mite life
cycle.
trochantin small, forward projecting sclerite at the base of the
trochanter.
trochus one of two ciliated rings of the typical rotiferan
corona; seecingulum.
troglobite obligate cave-dweller, often characterized in crustaceans by a reduction in eye structure and a lack of pigments; also, frequently demonstrating K-strategies.
troglophile organism living in a cave in a nonobligate relationship; such organisms usually complete their life history in the cave and commonly have conspecifics surviving outside the cave; contrasts with troglobite.
trophi complex set of hard jaws characteristically found in
the pharynx (mastax) of rotifers.
trophozoite mature, vegetative, feeding, adult stage in the
protozoan life cycle; used primarily with reference to
parasitic species of nonciliate taxa; synonymous with
trophont.
tubercles small, rounded knobs on the body surface of an animal, larger than papillae, smaller than nodes.
tun barrel-shaped form of a tardigrade in a cryptobiotic (latent) state.
turnover time amount of time necessary for the standing crop
biomass to be replaced completely in a species.
type taxon (or specimen) original individual of a species on
which the species name is based; type specimens are kept
in museums for comparative purposes and to reduce
confusion about scientific nomenclature.
typhlosole invaginated, cilated, particle-sorting structure running the length of the midgut and extending into the ciliated sorting surfaces of the floor of the stomach and digestive diverticulum in bivalve molluscs.
umbo in bivalve molluscs, portion of the shell raised above
the dorsal shell margin near the hinge (also called the
beak).
uncate condition of the pedipalp in deutonymphs and adults
of some mites in which the ventral surface of the tibia is
expanded and produced distally to oppose the tarsus,
forming an efficient grasping organ.
uncinate type of trophi (jaw structure) found in rotifers characterized by unci possessing few teeth, usually with one
large one and a few small ones.
uncus one of the seven pieces comprising rotifer trophi; an
uncus together with a manubrium form the malleus subunit.
undulating membrane compound ciliary apparatus typically
lying on the right side of the buccal cavity of some protozoa; synonomous with paroral membrane.
undulipodium swimming organelle of eukaryotic cells, including protozoa, includes both the flagellum and the cilium.
univoltine having one brood or generation per year.
urogomphus paired, unsegmented terminal appendage of
coleopteran larvae.
uroid posterior part of moving “naked” lobose amoeba.
urosome portion of the copepod body posterior to the major
body articulation.
Glossary
urstigma (urstigmata) knob-like structure with a porous cap
located between the first and second coxal plates in larval mites; the paired urstigmata are apparently homologous with the genital acetabula of deutonymphs and
adults and probably function as an osmoregulatory chloride epithelium.
uterus region in female reproductive system of invertebrates
that stores egg capsules prior to release.
vagina portion of the female reproductive tract connecting
the vulva with the uterus or uteri; a female tract used in
copulation, distal to entrance of the oviducts.
valve one of either the left or right shell of a seed shrimp
(Ostracoda) or mussel (Mollusca), generally referred to
as left or right valve. In nematodes, the disc or nipplelike structure which may occur in the metacorpus or
basal bulb of the pharynx.
varve layer of sediment representing one year’s accumulation.
vas deferens duct from the testes, through which spermatozoa
pass.
veliger free-swimming larval stage of molluscs characterized
by a ciliated velum used for swimming and feeding; in
North American freshwater bivalves, veligers occur only
in exotic Dreissena spp.
ventral shield single large sclerite or series of closely fitting
plates and platelets completely covering the venter of the
idiosoma in deutonymphs and adults of certain mite taxa.
verge penis or organ that bears the penis.
verticil setae most anterior series of sensory setae on the dorsal surface of the idiosoma, consisting of and internal
pair (vi) and an external pair (ve).
vesicula seminalis enlarged part of the sperm duct or ejaculatory duct holding sperm.
vestibule space between the outer lamella and the duplicature
in the proximal portion of an ostracode valve.
virgate type of trophi (jaw structure) found in rotifers that
are modified for piercing and pumping; generally can be
recognized by the long fulcrum and manubria; some are
asymmetrically shaped.
1019
vitellarium yolk gland associated with the ovary in bdelloid
and monogonont rotifers.
vitelline membrane most external membrane of a molluscan
egg.
viviparous type of reproduction in which the young develop
internally and are maintained and nourished by the
mother before birth; see oviparous and ovoviviparous.
VO2 abbreviation for volume of oxygen consumed per unit
time.
voltine refers to the number of generations during the year or
during the reproductive season (e.g., univoltine, bivoltine, polyvoltine).
vulva ventrally located genital opening of the female nematode, usually located in the midbody region.
water tube in bivalve molluscs, the channels formed between
the descending and ascending limbs of lamellibranch gill
filaments which carry water past gill ostia dorsally to the
epibranchial cavity; also form gill marsupial chambers in
which eggs and larval stages are incubated through development to juveniles or glochidia.
wetlands ecosystems whose soil is saturated for long periods
seasonally or continuously; include marshes, swamps,
ephemeral ponds, etc.
wing in bivalve molluscs, a thin, flattened, dorsally extended
portion of the posterior slope occurring on the shell of
some unionacean species.
YOY (young-of-the-year) juveniles during the period from
the last larval stage to adulthood, or one year of agewhichever comes sooner.
Zenker’s organ ejaculation ducts, composed of longitudinal
muscles in the shape of a tube on which occur radiating
bristles and are covered in a chitinous sheath in freshwater ostracodes.
zooid one of the physically connected, asexually replicated,
morphologic units that comprise a colony; in bryozoans, the zooid includes the polypide and associated
musculature as well as all parts of the adjacent body
wall.
This Page Intentionally Left Blank
Subject Index
Acanthametropodidae, see Ephemeroptera
Acanthobdellidae
anatomy
annuli, 468
digestive system, 469 – 470
excretory system, 471
eyes, 468
morphology, 467 – 468
papillae, 468
reproductive organs, 470
suckers, 468 – 469
vascular system, 471 – 472
behavior, 481
diagnostic features of leeches, 466
dispersal, 479 – 480
diversity, 474
ecology overview, 10
identification, 488
life cycle, 479
locomotion, 472
parasitism, 466 – 467,474
predation, 480 – 481
reproduction, 477 – 479
respiration, 472
taxonomic key, 489
taxonomic status, 466 – 467
Acariformes
Acaridida, 649
Actinedida, 648
Oribatida, 648 – 649
Acroloxidae, see Gastropoda
Adelphaga, see Coleoptera
Aeshinidae, see Odonata
Alderfly, see Megaloptera
Allen curve, secondary production
measurement, 758
Allozyme, variants in rotifer populations,
222 – 223
Ameletidae, see Ephemeroptera
Ametropodidae, see Ephemeroptera
Amphipoda, see Peracarida
Amphizoidae, see Coleoptera
Ampullaridae, see Gastropoda
Ancylidae, see Gastropoda
Anisoptera, see Odonata
Annelida, see also Acanthobdellidae;
Branchiobdellida; Euhirudinea;
Oligochaeta
controversy in taxonomy, 9 – 10,432,
467
Annuli
Acanthobdellidae, 468
Euhirudinea, 468
Annulipalpia, see Trichoptera
Anomopoda, see Cladocera
Anostraca, see Branchiopoda
Antennae
cladocera, 847,850
Crustacea, 776
Antennules
cladocera, 847,849 – 850
Crustacea, 776
Decapoda, 955
Apataniidae, see Trichoptera
Aquatic insect, see Insect, aquatic
Arachnida, see also Acariformes;
Dolomedes; Hydrachnida; Parisitiformes
classification, 549
1021
1022
Subject Index
Arthropleidae, see Ephemeroptera
Arthropoda, see also Crustacea; Hydrachnida; Insects, aquatic
aquatic insects, 12
crustaceans, 12 – 16
water mites, 12
Astacidae, see Decapoda
Athericidae, see Diptera
Atyidae, see Decapoda
Baetidae, see Ephemeroptera
Baetiscidae, see Ephemeroptera
Bdelloidea, see Rotifera
Behningiidae, see Ephemeroptera
Belostomatidae, see Heteroptera
Beraeidae, see Trichoptera
Bicarbonate, transport in bivalves, 334
Bicoecids, anatomy, 52
Binomial nomenclature, taxonomic classification, 2
Birth rate, equation, 221
Bithynidae, see Gastropoda
Bivalvia, see also Unionoidea
anatomy
gastric shield, 341
gills, 336 – 338
kidney, 339
lateral cilia, 337 – 338
locomotory structures, 335 – 336
mouth, 341
nervous system, 343 – 344
overview, 331
reproductive structures, 341 – 343
sensory organs, 344
shell, 333 – 335
stomach, 341
anoxia response, 349 – 350
behavioral ecology, 373,375 – 376
biomonitor applications, 390 – 391
brooding of embryos, 342
burrowing, 335 – 336,373
calcium requirements, 360
collection, 393 – 394
conservation biology, 357 – 358
culture, 395 – 397
desiccation resistance, 350 – 352
distribution, 332 – 333
dispersal, 354 – 356
pH effects, 360
stream size effects, 361 – 262
substrate effects, 358 – 360
temperature effects, 361
water depth effects, 360 – 361
diurnal cucles, 348
diversity, 332,354,356 – 358
economic importance, 9
ecosystem role, 387 – 390
evolution, 390 – 393
extinction, 356 – 357
feeding
filter feeding
particle capture, 376
rate of filtering, 381 – 382
sorting of filtered particles, 376,380
sediment feeding, 382 – 383
filtration rate, season effects, 347
glycogenic hormones, 3431
glycogen stores, season effects, 347 – 348
growth efficiency, 390
habitat, 9
hemolymph, 336
ion transport, 334,339
nitrogen excretion, 340,351
osmoregulation, 338 – 340
carbonic anhydrase role, 353
hemolymph osmolarity, 353
hormonal control, 354
ion transport, 352 – 353
salt tolerfance, 352
oxygen consumption
biomass relationship, 346
growth and reproduction relationship,
347
hypoxia response, 348 – 349
pollutant effects, 348
seasonal cycles, 344 – 346
population regulation
competitive interactions, 386 – 387
density-dependent mechanisms, 387
oxygen, 384
parasites, 384
pH, 383 – 384
pollution, 384
predation, 385 – 386
temperature, 383
water levels, 384
reproduction
egg, 343
sperm, 343
temperature regulation, 343
shell synthesis, 331,334 – 335
specimen preparation for identification,
394 – 395
subclasses, 331
taxonomic keys
Corbiculacea, 398 – 400
superfamilies, 397
Unionoidea, 400 – 403,408 – 415
Blephariceridae, see Diptera
Blood, cladocera, 854 – 855
Body wall, bryzoans, 507 – 508
Bosminidae, see Cladocera
Brachycentridae, see Trichoptera
Brachycera, see Diptera
Brain, see Nervous system
Branchiobdellida
anatomy
digestive system, 457
morphology, 456 – 457
nervous system, 457
reproductive organs, 457
vascular system, 457
collection, 459
distribution, 458
diversity, 456,458
evolution, 459
feeding, 457
habitat, 10
life cycle, 458
parasitism and parasites, 458 – 459
reproduction, 458
specimen preparation for identification,
459
taxonomic key, 459 – 460
Branchiopoda, see also Crustacea
adaptations of noncladocerans
diapause, 892
oxygen concentrations, 891
salinity, 891
temperature, 891
anatomic features
Anostraca, 887
Laevicaudata, 887
Notostraca, 887,889
Spinicaudata, 887
behavior
mating, 892 – 893
swimming, 892
cladocerans, see Cladocera
collection of noncladocerans, 895
conservation biology, 890 – 891
culture of noncladocerans, 895
distribution of noncladocerans, 890
ecosystem role of noncladocerans, 894
eggs on noncladocerans, 890
feeding mechanisms, 893 – 894
orders and families, 846
overview, 13 – 14
population regulation of noncladocerans,
894
prospects for research, 895
specimen preparation, 895
taxonomic key, 895,897,900
toxicology of noncladocerans, 894 – 895
Branchiura, see also Crustacea
distribution, 783 – 784
larva, 784
morphology, 784
overview, 13
reproduction, 784
taxonomic key, 795 – 796
Bryzoans, see also Entoprocta
anatomy
body wall, 507 – 508
coelum, 506
ectocyst, 508 – 509
lophophore, 505 – 506,509
nervous system, 506 – 507
statoblast, 506,510,517
zooid, 505 – 506
classifications, 505
collection, 516
culture, 516
distribution, 511 – 512
diversity, 511
environmental physiology, 510 – 511
evolution, 514 – 515
fecal deposits, 514
feeding
diet, 513
mechanism, 509
habitats, 505,511 – 512
larvae, 509
life cycle, 512
Subject Index
overview, 10 – 11
parasites, 513
predation, 513
reproduction
asexual budding, 509 – 510
fission, 510
gametogenesis, 513
sexual, 509
specimen preparation for identification,
516 – 517
taxonomic key, 517 – 523
Caddisfly, see Trichoptera
Caenidae, see Ephemeroptera
Calamoceratidae, see Trichoptera
Calanoida, see Copepoda
Calcium
carbonate shell synthesis in bivalves,
334 – 335
composition of bivalve shells, 333 – 334
crustacean requirements, 781
Gastropoda requirements, 302
mussel requirements, 27,360
transport in bivalves, 334
Calopterygidae, see Odonata
Cambaridae, see Decapoda
Capniidae, see Plecoptera
Carapace, cladocera, 850
Caterpillar, see Lepidoptera
Caves, see Underground aquatic habitats
Ceratopogonidae, see Diptera
Chaetae, Oligochaeta, 435 – 436,443
Chaoboridae, see Diptera
Chironomidae, see Diptera
Chloroperlidae, see Plecoptera
Choanocyte, sponges, 101
Choanoflagellates, anatomy, 51 – 52
Chrysomellidae, see Coleoptera
Chrysophytes, anatomy, 52
Chydoridae, see Cladocera
Cilia
bivalves, 337 – 338
Protozoa, 47 – 48
Ciliophora, anatomy, 54 – 55
Circadian clock, Decapoda, 959
Circulatory system
cladoceran heart, 851
Copepoda, 914
Crustacea, 777
Decapoda, 954
Gastropoda, 301
Ostracoda, 812
Cladistics, taxonomic classification, 1
Cladocera, see also Crustacea
anatomy
antennae, 847,850
antennules, 847,849 – 850
carapace, 850
digestive system, 851
eye, 847,849
fat globules, 851
head, 847,850
heart, 851
legs, 850 – 851
maxillae, 850
morphology, 847
reproductive organs, 852
behavior
diel vertical migration, 860 – 861
escape, 861 – 862
mating, 859 – 860
swarming, 861
swimming, 860
blood, 854 – 855
collection, 869 – 870
coloration, 850
conservation biology, 858
culture, 870 – 871
cyclomorphosis, 851 – 852
distribution, 856 – 858
ecosystem role, 864
eggs, 853 – 854
feeding
herbivores, 862
mechanism, 850 – 851
predators, 862
fossil record, 865,867
life cycle, 852 – 854
metabolic rate, factors affecting
body size, 855
food concentration, 855
interaction between factors, 855 – 856
temperature, 855
molting, 854
orders and families, 846 – 847
overview, 13 – 14
paleolimnology applications, 864 – 865
phylogenetic analysis, 867
population regulation, 863 – 864
predation, 863 – 864
prospects for research, 869
reproduction, 852 – 854
species distinction, 856
species diversity effects
competitive interactions, 857 – 858
lake size, 857
littoral species, 858
pH, 857
salinity, 857
turbidity, 857
specimen preparation, 871
taxonomic keys
Bosminidae, 881
Chydoridae, 874 – 878
Daphniidae, 879 – 881
families, 871,873
Halopediidae, 873
Haplopoda, 885,887
Macrothricoidea, 883
Moinidae, 879 – 881
Onychopoda, 885,887
Sididae, 873 – 874
toxicology, 867,869
Claws, Tardigrada, 530 – 532
Clearance rate, calculation, 925 – 926
Clitellata
controversy in taxonomy, 432,466 – 467
evolution, 441 – 442,466
Clitellum, Oligochaeta, 432,434
Cnidaria,
anatomy
body plan, 136 – 137
nematocysts, 137 – 138,140
overview, 6,136
classification, 151
collection, 150
culture, 150 – 151
ecological interactions, 140
feeding, 138
molecular biology, 145 – 146
reproduction, 138 – 140
taxa, 135 – 136
taxonomic key, 152 – 153
Coelum, bryzoans, 506
Coenagrionidae, see Odonata
Coleoptera, see also Insect, aquatic
Adelphaga
Amphizoidae, 684
Dytiscidae, 684 – 685
Gyrinidae, 685
Haliplidae, 685
Noteridae, 685 – 686
overview, 684
aquatic species, 682
collection, 682
literature references to species keys,
688 – 689
morphology
adults, 682
larvae, 683
Myxophaga, Hydroscaphidae, 686
Polyphaga
Chrysomellidae, 687
Curculionidae, 687
Dryopidae, 686 – 687
Elmidae, 687
Helophoridae, 688
Hydraenidae, 687 – 688
Hydrochidae, 688
Hydrophilidae, 688
Lampyridae, 688
Lutrochidae, 687
overview, 686
Psephenidae, 687
Ptilodactylidae, 687
Scirtidae, 688
respiration, 682 – 683
taxonomic keys
adults, 708,710 – 711
larvae, 712 – 714
Collection
aquatic insects, 659
Branchiobdellida, 459
bryzoans, 516
cladocera, 869 – 870
Cnidaria, 150
Coleoptera, 682
Collembola, 696
Copepoda, 930
Crustacea, 794 – 795
Decapoda
crayfish, 985 – 986
shrimp
Atyidae, 985
1023
1024
Subject Index
Collection (continued)
Palaemonidae, 985
Diptera, 689
Euhirudinea, 486
Gastropoda, 315
Gastrotricha, 188
Hydrachnida
deutonymphs and adults, 583 – 584
larvae, 584 – 585
Nematoda
extraction, 265 – 266,268
sampling, 265
Nematomorpha, 290 – 291
Nemertea, 176
noncladocerans, 895
Oligochaeta, 444
Ostracoda, 825 – 826
Porifera, 115
Protozoa, 69 – 70
Rotifera, 230 – 231
Tardigrada, 541
Trichoptera larvae, 671 – 672
Turbellaria, 163
Collembola
classification, 695
collection, 696
life cycle, 695 – 696
morphology, 695
overview, 12
specimen preparation, 696
taxonomic key, 716 – 717
Colpodea, anatomy, 55
Conservation biology
aquatic insects
biomonitoring studies, 763 – 764
disturbance effects
flow impairment, 761 – 762
insecticide spills, 762 – 763
natural versus human disturbances,
761
land use effects on communities
agriculture, 761
large woody debris alterations, 760
lentic environments, 759
road construction, 761
streams, 759 – 760
wetlands, 759
Bivalvia
biomonitor uses, 390 – 391
conservation biology, 357 – 358
Branchiopoda, 890 – 891
cladocera, 858
Crustacea water quality response and biomonitor use, 790
Decapoda, 982 – 984
Euhirudinea as bioindicator, 485 – 486
Gastropoda, 315
Hydrachnida as bioindicator, 582 – 583
Plecoptera biomonitoring studies,
763 – 764
Trichoptera biomonitoring studies,
763 – 764
Unionoidea, 368
Consumption, equation, 263
Contractile vacuole, Protozoa, 46 – 47
Copepoda, see also Crustacea
adaptations
oxygen concentration, 921 – 922,927
temperature, 921
anatomy
circulatory system, 914
digestive system, 914
head, 912
legs, 912,914
morphology, 911 – 912,914
nervous system, 916
reproductive organs, 914,916
sensory organs, 916
sex differences, 917
thorax, 912
behavior
feeding, 92
mating, 919
swimming, 922 – 923
classification, 911 – 912
collection, 930
culture, 931
diapause, 917,919
dissection techniques for identification,
931 – 932
distribution, 920 – 921
diversity, 911
ecosystem role, 928 – 929
eggs, 917
feeding
diet, 923 – 924
mechanisms, 924 – 925
mosquito control, 929
rate calculations, 925 – 926
habitats, 912
identification, 932 – 934
life cycle, 916 – 917,919 – 920
nitrogen excretion, 914
overview, 14 – 15
parasitism, 929
population regulation
birth rate, 928
competition, 927
death rate, 928
food limitation effects, 920,927
growth rates, 926,928 – 929
predation, 920,927
temperature effects, 926 – 927
prospects for research, 929 – 930
reproduction, 916 – 917,919
sexing, 932 – 933
species
Calanoida, 948
Cyclopoida, 949
Harpacticoida, 950
Poecilostomatoida, 950
specimen preparation, 930 – 931
staging, 933
taxonomic keys
Calanoida, 934 – 935
Cyclopoida, 936 – 937
Harpacticoida, 937 – 938
behavioral ecology, 373,375
collection, 394
competitive interactions, 386
culture, 395 – 397
density-dependent mechanisms of population regulation, 387
ecosystem role, 388 – 390
environmental stresses, 371
evolution, 392
life cycle, 370 – 372
postreproductive die-offs, 384
reproduction, 370 – 371
specimen preparation for identification,
394 – 395
taxonomic key for Corbiculacea,
398 – 400
Cordulegastridae, see Odonata
Corduliidae, see Odonata
Corethrellidae, see Diptera
Corixidae, see Heteroptera
Corona, Rotifera, 198 – 199,201
Corydalidae, see Megaloptera
distribution, 146 – 147
feeding, 147
growth, 147
life cycle, 147 – 148
morphology, 146
reproduction, 147
Crayfish, see Decapoda
Cryptobiosis, Tardigrada
encystment, 535 – 536
anoxybiosis, 536
cryobiosis, 536
osmobiosis, 536
anhydrobiosis, 536
Crustacea, see also Branchiopoda;
Branchiura; Cladocera; Copepoda;
Decapoda; Ostracoda; Peracarida
anatomy
antennae, 776
antennules, 776
appendages, 776
circulatory system, 777
digestive system, 776 – 777
maxillae, 776
maxillules, 776
morphology, 776
nervous system, 779
reproductive organs, 779 – 780
respiratory system, 777
skeleton, 776
calcium requirements, 781
chemoreception, 782
classification, 773 – 774
collection, 794 – 795
culture, 795
distribution
ancient lake studies, 794
dispersal barriers, 793 – 794
groundwaters, 792 – 793
historical perspective, 791 – 792
surface waters, 793
ecological impact, 773 – 774
embryogeny, 780
evolution, 773,783
feeding
mechanism, 776 – 777
resources, 788 – 789
Subject Index
vertical migration, 789 – 790
fossil record, 773
habitats, 780,787 – 788
longevity, 790 – 791
mechanoreception, 782
molting, 780
mysid predation, 789 – 790
nitrogen excretion, 777
osmoregulation, 777,779
photoreception, 781 – 782
population genetics, 793 – 794
specimen preparation, 795
taxonomic key to orders, 795 – 796
thermoreception, 783
tolerances
pH, 781
salt, 780 – 781
temperature, 781
water quality response and biomonitor
use, 790
Cryptomonads, anatomy, 52
Ctenopoda, see Cladocera
Culicidae, see Diptera
Culture
aquatic insects, 659 – 660
Bivalvia, 395 – 397
bryzoans, 516
cladocera, 870 – 871
Cnidaria, 150 – 151
Copepoda, 931
Crustacea, 795
Decapoda, 986
Euhirudinea, 486
Gastropoda, 315
Gastrotricha, 188 – 189
Hydrachnida, 585
Nematoda, 269
Nemertea, 176
noncladocerans, 895
Oligochaeta, 444 – 445
Ostracoda, 826 – 827
Porifera, 115
Protozoa, 69 – 70
Rotifera, 231
Turbellaria, 163
Curculionidae, see Coleoptera
Cuticle
Nematoda, 256,258
Nematomorpha, 282 – 283
Tardigrada, 528 – 530
Cyclomorphosis, cladocera, 851 – 852
Cyclopoida, see Copepoda
Damselfly, see Odonata
Daphniidae, see Cladocera
Death rate, equation, 222
Decapoda
adaptations
coloration, 959
temperature, 958
anatomy
antenulles, 955
biramous appendages, 952
circulatory system, 954
digestive system, 952,954
eye, 955
gills, 954
morphology, 951 – 952
nervous system, 954 – 955
sensory organs, 955
statocyst, 955
behavior of crayfish
burrowing, 974,976 – 977
pH effects, 977 – 978
predation avoidance, 978 – 980
shelter and aggression, 978
water current effects, 977
circadian clock, 959
collection
crayfish, 985 – 986
shrimp
Atyidae, 985
Palaemonidae, 985
conservation biology, 982 – 984
culture, 986
distribution
crayfish
Astacidae, 966 – 967
Cambaridae, 967 – 968
overview, 963,965 – 966
shrimp
Atyidae, 960 – 961
Palaemonidae, 961 – 963
diversity, 774,960
economic impact of crayfish, 974,985
ecosystem role, 973 – 974
feeding
crayfish
behavior, 975 – 976
chemoreception, 976
diet, 974 – 975
predation, 975
shrimp, 974
genera and subgenera, 963 – 964
habitats, 973 – 974
life cycle
crayfish, 969 – 970,972 – 973
shrimp, 968 – 969
molting, 957 – 958
osmoregulation, 959
overview, 15 – 16
parasitism, 974
population regulation
crayfish factors
competition, 981
displacent by invaders, 982
growth rates, 981
hybridization, 981 – 982
overview, 980
predation, 980 – 981
shrimp, 980
prospects for research, 982 – 985
reproduction
crayfish, 969 – 970,972 – 973
gonads, 958
pheromones, 958,983
shrimp, 968 – 969
respiration, 955 – 957
specimen preparation, 986 – 987
1025
taxonomic key, 987 – 989
toxicology, 959 – 960
Detrivory
aquatic insects
coarse particulate organic matter processing, 747
DOM ingestion, 748
fine particulate organic matter formation, 747 – 748
large woody debris, 748
Deuterophlebiidae, see Diptera
Diapause
Copepoda, 917,919
noncladoceran adaptation, 892
Digestion system
Acanthobdellidae, 469 – 470
Branchiobdellida, 457
cladocera, 851
Copepoda, 914
Crustacea, 776 – 777
Decapoda, 952,954
Euhirudinea, 469 – 470
Gastropoda, 300 – 301
Gastrotricha, 182
Hydrachnida, 567
Nematomorpha, 283
Ostracoda, 812
Porifera digestion, 102
Tardigrada
buccal aparatus, 532 – 534
esophagus, 534
hindgut, 534
midgut, 534
overview, 532
Dinoflagellates, anatomy, 52
Diplomonads, anatomy, 52
Dipseudopsidae, see Trichoptera
Diptera, see also Insect, aquatic
Brachycera
Athericidae, 693
Dolicopodidae, 693
Empididae, 693 – 694
Ephydridae, 694
Muscidae, 694
Sciomyzidae, 694
Stratiomyidae, 694
Syrphidae, 694 – 695
Tabanidae, 695
collection, 689
habitats, 688
larva, 689,693
literature references to species keys, 690
Nematocera
Blephariceridae, 690
Ceratopogonidae, 690
Chaoboridae, 690 – 691
Chironomidae, 691
Corethrellidae, 691
Culicidae, 691 – 692
Deuterophlebiidae, 692
Dixidae, 692
Nymphomyiidae, 692
Psychodidae, 692
Ptychopteridae, 692
Simuliidae, 692
1026
Subject Index
Diptera (continued)
Tanyderidae, 692 – 693
Thaumaleidae, 693
Tipulidae, 693
pupa, 689
taxonomic key to larvae, 714 – 716
Discobolocyst, Protozoa, 51
Distribution
Bivalvia, 332 – 333
dispersal, 354 – 356
pH effects, 360
stream size effects, 361 – 262
substrate effects, 358 – 360
temperature effects, 361
water depth effects, 360 – 361
Branchiobdellida, 458
Branchiura, 783 – 784
bryzoans, 511 – 512
cladocera, 856 – 858
Copepoda, 920 – 921
Crustacea
ancient lake studies, 794
dispersal barriers, 793 – 794
groundwaters, 792 – 793
historical perspective of study,
791 – 792
surface waters, 793
Decapoda
crayfish
Astacidae, 966 – 967
Cambaridae, 967 – 968
overview, 963,965 – 966
shrimp
Atyidae, 960 – 961
Palaemonidae, 961 – 963
Gastropoda
Acroloxidae, 302 – 303
Ampullaridae, 302 – 303
Ancylidae, 302
Bithynidae, 302
ecological determinants, 309 – 312
Hydrobiidae, 302 – 303
Lymnaeidae, 302 – 303
Micromelaniidae, 302
Neritinidae, 302 – 303
Physidae, 302 – 304
Planorbidae, 302,304
Pleuroceridae, 302
Pomatiopsidae, 302 – 303
Thiaridae, 302
Valvatidae, 302
Viviparidae, 302 – 303
Gastrotricha, 181,183 – 185
Hydrachnida, 552
Nemertea, 175
Oligochaeta, 438 – 439
Ostracoda
benthic nonswimmers, 816
benthic swimmers, 816
Porifera, 106 – 107,117 – 127
Protozoa, 59
Rotifera, 195 – 196,208
Tardigrada, 540
Turbellaria, 156,159 – 160,162
Unionoidea, 402
Diversity
Acanthobdellidae, 474
aquatic insects, 729 – 730
Bivalvia, 332,354,356 – 358
Branchiobdellida, 456,458
bryzoans, 511
Copepoda, 911
Decapoda, 774,960
Euhirudinea, 474
Gastropoda, 297,302
Gastrotricha, 183
Hydrachnida, 549 – 551
Oligochaeta, 438 – 439
Porifera, 105 – 106
Protozoa, 58 – 59
Trichoptera, 670
Dixidae, see Diptera
Dolicopodidae, see Diptera
Dragonfly, see Odonata
Dryopidae, see Coleoptera
Dytiscidae, see Coleoptera
Earthworm, see Oligochaeta
Ecdysis, see Molting
Ecdysteroids, molting stimulation in
crayfish, 958
Ecnomidae, see Trichoptera
Ecosystem role
Bivalvia, 387 – 390
Branchiopoda
cladocerans, 864
noncladocerans, 89
Copepoda, 928 – 929
Corbicula fluminea, 388 – 390
Decapoda, 973 – 974
Nemertea, 175 – 176
Odonata, 665
Protozoa, 4 – 5,43
Rotifera, 224 – 225
Trichoptera, 672
Turbellaria, 161 – 162
Unionoidea, 387
Ectocyst, bryzoans, 508 – 509
Egg, see also Reproduction
cladocera, 853 – 854
Copepoda, 917
Hydrachnida, 568 – 569
Ostracoda, 813,822
Ejectisome, Protozoa, 48,51
Elmidae, see Coleoptera
Empididae, see Diptera
Entoprocta, see also Bryzoans
anatomy, 515 – 516
distribution, 516
evolution, 516
feeding, 516
species, 515
Ephemerellidae, see Ephemeroptera
Ephemeridae, see Ephemeroptera
Ephemeroptera, see also Insect, aquatic
biomonitoring studies, 763 – 764
feeding, 661
identification, 661
life cycle, 660 – 661
literature references to species keys, 662
Pisciforma
Acanthametropodidae, 661 – 662
Ameletidae, 662
Ametropodidae, 662
Baetidae, 662
Metretopodidae, 662 – 663
Pseudironidae, 663
Siphlonuridae, 663
reproduction, 661
Retrachaeta
Baetiscidae, 663
Behningiidae, 663
Caenidae, 663
Ephemerellidae, 663
Ephemeridae, 663 – 664
Leptohyphidae, 664
Leptophlebiidae, 664
Neoephemeridae, 664
Polymitarcyidae, 664
Potamanthidae, 664
Setisura
Arthropleidae, 664
Heptageniidae, 664
Isonychiidae, 664 – 665
Oligoneuriidae, 665
taxonomic key to larvae, 697 – 700
Ephydridae, see Diptera
Erpobdellidae, see Euhirudinea
Euglenids, anatomy, 52
Euhirudinea
anatomy
annuli, 468
digestive system, 469 – 470
excretory system, 471
eyes, 468
morphology, 468
papillae, 468
reproductive organs, 470
suckers, 468 – 469,475
vascular system, 471 – 472
collection, 486
culture, 486
dispersal, 479 – 480
diversity, 474
ecology overview, 10
feeding, 465 – 466,474 – 477
foraging behavior
oxygen level effects, 483
predators, 481 – 483
sanguivores, 481
hyperoxia tolerance, 473
hypoxia tolerance, 472 – 473
identification
diagnostic features of leeches,
466,487 – 488
Erpobdellidae, 489
Glossiphoniidae, 488
Hirudinidae, 489
Piscicolidae, 488 – 489
specimen preparation for identification,
486 – 487
life cycle, 478 – 479
locomotion, 472
population regulation, 483,485
Subject Index
predation, 480 – 481
reproduction, 471,477 – 479
respiration, 472
salt tolerance, 473 – 474
taxonomic key, 489 – 497
taxonomic status, 466 – 467
toxicology and use as bioindicator,
485 – 486
Euholognatha, see Plecoptera
Evolution
Bivalvia, 390 – 393
Branchiobdellida, 459
bryzoans, 514 – 515
cladocera, 865,867
Clitellata, 441 – 442,466
Crustacea, 773,783
Gastropoda
phylogenetic trees, 313 – 314
pulmonates, 313
ribosomal DNA analysis, 313 – 314
Gastrotricha, 187 – 188
Hydrachnida
exoskeleton, 564,566 – 567
overview, 550
Nematoda, 261 – 262
Nematomorpha, 280 – 282
Oligochaeta
clitellate taxa, 443 – 444
Enchytraeidae, 441
Haplotaxidae, 441
Lumbriculidae, 443
Naididae, 441,443
phylogenetic tree of Annelida, 442
Tubificidae, 441,443
Ostracoda, 807 – 808,827
Peracarida, 785 – 786
Porifera, 114 – 115
Protozoa, 68 – 69
Sphaeriidae, 390,392 – 393
Tardigrada, 527
Unionoidea, 390 – 393
Excretory system
Acanthobdellidae, 471
Euhirudinea, 471
Gastropoda, 301
Gastrotricha, 182 – 183
Hydrachnida, 567
Nematoda, 259
Tardigrada, 535
Extrusome, Protozoa, 48,51
Eye
Acanthobdellidae, 468
cladocera, 847,849
Decapoda, 955
Euhirudinea, 468
Oligochaeta spots, 437 – 438
Ostracoda, 812
Turbellaria, 157
Fat globules, cladocera, 851
Feeding, see also Detrivory; Grazing; Predation
Bivalvia
filter feeding
particle capture, 376
rate of filtering, 381 – 382
sorting of filtered particles, 376,380
sediment feeding, 382 – 383
Branchiobdellida, 457
bryzoans
diet, 513
mechanism, 509
cladocerans
herbivores, 862
mechanism, 850 – 851
predators, 862
Copepoda
diet, 923 – 924
mechanisms, 924 – 925
mosquito control, 929
rate calculations, 925 – 926
Crustacea
mechanism, 776 – 777
resources, 788 – 789
vertical migration, 789 – 790
Decapoda
crayfish
behavior, 975 – 976
chemoreception, 976
diet, 974 – 975
predation, 975
shrimp, 974
Ephemeroptera, 661
Euhirudinea,
465 – 466,474 – 477,481 – 483
Gastropoda, 305 – 307
Gastrotricha, 186 – 187
Hydrachnida
diet, 581
foraging behavior, 581 – 582
prey population effects, 581
Nematoda, 261 – 263
Nemertea, 174
noncladoceran mechanisms, 893 – 894
Oligochaeta, 437,440
Ostracoda, 823 – 824
Peracarida, 788 – 790
Plecoptera, 668 – 669
Porifera, 110 – 112
Rotifera, 223 – 224
Turbellaria, 160 – 161
Filosea, anatomy, 54
Filtering rate
bivalves, 381 – 382
equation, 381
Fishfly, see Megaloptera
Flagella, Protozoa, 47 – 48
Flatworms, see Nemertea; Turbellaria
Flowing waters, see Lotic environments
Food vacuole, Protozoa, 45 – 46
Foot
Gastropoda, 299
Rotifera, 197
Gastric shield, Bivalvia, 341
Gastropoda
anatomy
circulatory system, 301
1027
digestive system, 300 – 301
excretory system, 301
foot, 299
head, 299
overview, 297
reproductive organs, 299 – 300
respiratory system, 301
shell, 298 – 299
visceral mass, 299
calcium requirements, 302
collection, 315
conservation biology, 315
culture, 315
dispersal ability, 310
distribution
Acroloxidae, 302 – 303
Ampullaridae, 302 – 303
Ancylidae, 302
Bithynidae, 302
ecological determinants, 309 – 312
Hydrobiidae, 302 – 303
Lymnaeidae, 302 – 303
Micromelaniidae, 302
Neritinidae, 302 – 303
Physidae, 302 – 304
Planorbidae, 302,304
Pleuroceridae, 302
Pomatiopsidae, 302 – 303
Thiaridae, 302
Valvatidae, 302
Viviparidae, 302 – 303
diversity, 297,302
evolution, 298
evolution
phylogenetic trees, 313 – 314
pulmonates, 313
ribosomal DNA analysis, 313 – 314
feeding, 305 – 307
habitat, 8 – 9,305
hypoxia adaptation, 302
life cycle, 304 – 305
pH effects, 309
population size factors
nutrition, 307 – 308
trematode parasites, 308 – 309
predation, 297,310 – 312
production ecology, 309
prospects for ecology research, 312
reproduction, 299 – 300,304 – 305
specimen preparation for identification,
315 – 316
subclasses, 297 – 298
taxonomic key, 316,318,320,323
temperature effects, 301 – 302
Gastrotricha
anatomy
brain, 182
digestive system, 182
excretory system, 182 – 183
morphology, 182
reproductive organs, 182
classification, 181 – 182
collection, 188
culture, 188 – 189
density and biomass, 183 – 185,187
1028
Subject Index
Gastrotricha (continued)
distribution, 181,183 – 185
diversity, 183
evolution, 187 – 188
feeding, 186 – 187
growth, 187
habitat, 7,183 – 184
life cycle, 185 – 186
predation, 187
preservation of specimens, 188
reproduction, 185 – 186
taxonomic key, 189 – 191
Gemmosclere, sponge spicule,
103 – 104,115 – 116
Gerridae, see Heteroptera
Gerromorpha, see Heteroptera
Gills
Bivalvia, 336 – 338
Decapoda, 954
Oligochaeta, 436
Unionoidea calcium phosphate concretions, 350,352
Glochidium, Unionoidea
collection, 394
fecindity, 364,366
fish parasitism, 363 – 364,368
forms, 363
host food mimicry, 364
host reduction effects, 385
size versus number released, 366
Glossiphoniidae, see Euhirudinea
Glossosomatidae, see Trichoptera
Goeridae, see Trichoptera
Gomphidae, see Odonata
Granuloreticulosida, anatomy, 54
Grazing, aquatic insects
algae crop, 749 – 750
ecosystem impact, 748 – 749
host-specific associations, 750,752
macrophyte grazing, 752
species distribution, 749
Growth efficiency, percent net growth efficiency equation, 390
Growth rate, equation for populations, 221
Gyrinidae, see Coleoptera
Habitat
aquatic insects
lentic communities
littoral communities, 734 – 735
profundal communities, 735
wetland communities, 735 – 736
lotic communities
drift of insects, 741 – 742
flow effects, 737 – 738
functional feeding groups, 738 – 740
recolonization mechanisms,
738 – 739
river continuum concept, 740 – 741
substrates, 736 – 737
saline habitats, 742
specialized habitats, 743
Bivalvia, 9
bryzoans, 505,511 – 512
Copepoda, 912
Crustacea, 780,787 – 788
Decapoda, 973 – 974
Diptera, 688
flowing waters, see Lotic environments
Gastropoda, 8 – 9,305
Gastrotricha, 7,183 – 184
Hydrachnida
interstitial habitats, 578 – 579
lakes, 579
riffle habitats, 578
springs, 578
stenothermic pools, 579
temporary pools, 579
inland water percentage of biospheric water, 19
lentic habitats, see Lentic ecosystems
Nematoda, 262 – 265
Nemertea, 7,175
Ostracoda
bromeliads, 821
carbonate content, 816,819
chemical water types, 817
diversity, 816
lakes, 821
oxygen concentrations, 818 – 819
pH tolerance, 818
population regulation, 824 – 825
salinity, 817 – 818
sediment – water interface, 822
solute composition, 818
streams, 821
temperature, 819 – 821
underground waters, 821 – 822
Sphaeriidae, 369 – 370
Tardigrada, 537 – 539
Turbellaria, 7
underground waters, see Underground
aquatic habitats
Hairworm, see Nematomorpha
Haliplidae, see Coleoptera
Halopediidae, see Cladocera
Haplopoda, see Cladocera
Haptocyst, Protozoa, 51
Harpacticoida, see Copepoda
Hatching
Nematoda, 260 – 261
Ostracoda, 814 – 815
Hatching rate, equation, 221
Head
cladocera, 847,850
Copepoda, 912
Gastropoda, 299
Ostracoda, 809,811
Heart, see Circulatory system
Hebridae, see Heteroptera
Helicopsychidae, see Trichoptera
Heliozoa, anatomy, 54
Helophoridae, see Coleoptera
Hemolymph
bivalves, 336,353
crustaceans, 777
Heptageniidae, see Ephemeroptera
Heteroptera, see also Insect, aquatic
aquatic and semiaquatic species, 677
Gerromorpha
Gerridae, 678 – 679
Hebridae, 679
Hydrometridae, 679
Mesoveliidae, 679
overview, 678
Veliidae, 679
identification, 677 – 678
literature references to species keys, 678
Nepomorpha
Belostomatidae, 680
Corixidae, 680
Naucoridae, 680
Nepidae, 680
Notonectidae, 681
overview, 679 – 680
Pleidae, 681
taxonomic key, 708
Hirudinidae, see Euhirudinea
Hydra, see also Cnidaria
distribution, 141
feeding, 142 – 143
growth, 142,144
habitat, 141
molecular biology, 145 – 146
morphology, 140,145
regeneration and immortality, 143,145
reproduction, 141 – 142
species groups, 151 – 152
substrates, 140 – 141
symbiosis with algae, 143 – 145
toxicology, 145
Hydrachnida
anatomy
digestive system, 567
excretory system, 567
morphology
adults, 560,563 – 564
larva morphology, 560 – 561
nervous system, 567 – 568
reproductive organs, 568
respiratory system, 567
collection
deutonymphs and adults, 583 – 584
larvae, 584 – 585
culture, 585
deutonymph stage, 575
diversity, 549 – 551
drawings
adult, 605 – 628
larva, 561,562,563,565,571,586 – 604
eggs, 568 – 569
evolution
exoskeleton, 564,566 – 567
overview, 550
exoskeleton sclerotization, 576
feeding
diet, 581
foraging behavior, 581 – 582
prey population effects, 581
global distribution, 552
habitats
interstitial habitats, 578 – 579
lakes, 579
riffle habitats, 578
Subject Index
springs, 578
stenothermic pools, 579
temporary pools, 579
life cycle, overview, 568
origins
Gondwana, 552
Laurasia, 552
North America, 552,560
Pangea, 552
parasitism of insects by larva
attachment and engorgement, 574 – 575
detachment, 575
ecological impact
host populations, 580 – 581
individual hosts, 580
intraspecific effects on same host,
581
emergence of larvae, 572 – 573
host selection, 569 – 570
infection, 574
recognition of host, 573
seeking and locating hosts, 573
site selection, 570 – 572
success rates, 569
population density, 550
predation, 582
protonymph stage, 575
reproduction
pheromones, 577
sex ratios, 578
spermatophore transfer
association during transfer, 576 – 577
copulation during transfer,
577 – 578
dissociation during transfer, 576
scanning electron micrographs,
629 – 630,632
specimen preparation, 585
taxonomic keys
adults, 638 – 648
larvae, 633 – 638
overview, 585 – 586
taxonomic relationships, 550 – 552
toxicology and use as bioindicator,
582 – 583
tritonymph stage, 576
Hydraenidae, see Coleoptera
Hydrobiidae, see Gastropoda
Hydrobiosidae, see Trichoptera
Hydrochidae, see Coleoptera
Hydrometridae, see Heteroptera
Hydrophilidae, see Coleoptera
Hydropsychidae, see Trichoptera
Hydroptilidae, see Trichoptera
Hydroscaphidae, see Coleoptera
Hypodermis, Nematoda, 258
ICZN, see International Code of Zoological
Nomenclature
Identification, see Specimen preparation;
Taxonomic keys
Increment-summation method, secondary
production measurement, 758
Ingestion rate, calculation, 925
Insect, aquatic
biomonitoring studies, 763 – 764
collection, 659
culture, 659 – 660
detrivory
coarse particulate organic matter processing, 747
detritus fractions, 747
DOM ingestion, 748
fine particulate organic matter formation, 747 – 748
large woody debris, 748
disturbance effects
flow impairment, 761 – 762
insecticide spills, 762 – 763
natural versus human disturbances,
761
diversity, 729 – 730
food web
gut analysis, 755 – 756
insects as energy flow conduits,
754 – 755
stable isotope studies, 756 – 757
trophic levels, 755
grazing
algae crop, 749 – 750
ecosystem impact, 748 – 749
host-specific associations, 750,752
macrophyte grazing, 752
species distribution, 749
land use effects on communities
agriculture, 761
large woody debris alterations, 760
lentic environments, 759
road construction, 761
streams, 759 – 760
wetlands, 759
lentic communities
littoral communities, 734 – 735
profundal communities, 735
wetland communities, 735 – 736
life cycles, 657 – 658
diversity, 743 – 744
hemimetabolous versus
holometabolous development,
746 – 747
synchronized development, 744,746
voltinism, 744,746
lotic communities
drift of insects, 741 – 742
flow effects, 737 – 738
functional feeding groups, 738 – 740
recolonization mechanisms, 738 – 739
river continuum concept, 740 – 741
substrates, 736 – 737
morphology
abdomen, 659
head, 658
thorax, 658 – 659
orders, see Coleoptera; Diptera;
Ephemeroptera; Heteroptera;
Lepidoptera; Megaloptera;
Neuroptera; Odonata; Plecoptera;
Trichoptera
physical constraints
1029
oxygen concentrations in water,
731 – 732
pH, 733
salt tolerance, 733 – 734
substrate and flow, 734
temperature, 732 – 733
predator – prey interactions
fish predation, 753 – 754
insects as predators, 752 – 754
lentic versus lotic environments,
752 – 754
saline habitats, 742
secondary production, 757 – 758
specialized habitats, 743
specimen preparation, 660
taxomic key to orders, 696 – 697
Instantaneous growth method, secondary
production measurement, 758
Integripalpia, see Trichoptera
International Code of Zoological Nomenclature (ICZN), 2
Internet resources, freshwater invertebrate
taxonomy, 3 – 4
Intestine, Nematoda, 258 – 259
Isonychiidae, see Ephemeroptera
Isopoda, see Peracarida
Jellyfish, see Craspedacusta
Kidney, Bivalvia, 339
Kinetoplastids, anatomy, 52
Laevicaudata, see Branchiopoda
Lakes, see Lentic ecosystems
Lampyridae, see Coleoptera
Land use, effects on aquatic insect communities
agriculture, 761
large woody debris alterations, 760
lentic environments, 759
road construction, 761
streams, 759 – 760
wetlands, 759
Larva
Branchiura, 784
bryzoans, 509
Coleoptera, 683
Diptera, 689,693
Hydrachnida, 560 – 561
Nematomorpha, 284
Odonata, 665 – 666
Plecoptera, 668
Trichoptera
adaptations, 671
collection, 671 – 672
pupate, 671
Leech, see Euhirudinea
Legs
cladocera, 850 – 851
Copepoda, 912,914
Ostracoda, 812
1030
Subject Index
Lentic ecosystems
ephemeral ponds, 36
insect communities
littoral communities, 734 – 735
profundal communities, 735
wetland communities, 735 – 736
lakes
abiotic zonation, 33 – 34
biotic zonation
littoral and profundal benthos,
35 – 36
neuston, 34 – 35
overview, 34
zooplankton of pelagic and littoral
habitats, 35
geomorphology, 33
saline lakes
abiotic characteristics, 38
geographic distribution, 38
species diversity, 38 – 39
swamps, 36,38
wetlands, 36
Lepidoptera, see also Insect, aquatic
larva, 681 – 682
morphology, 682
Lepidostomatidae, see Trichoptera
Leptoceridae, see Trichoptera
Leptohyphidae, see Ephemeroptera
Leptophlebiidae, see Ephemeroptera
Lestidae, see Odonata
Libellulidae, see Odonata
Life cycle
Acanthobdellidae, 479
aquatic insects
diversity, 743 – 744
hemimetabolous versus
holometabolous development,
746 – 747
synchronized development, 744,746
voltinism, 744,746
Branchiobdellida, 458
bryzoans, 512
cladocera, 852 – 854
Collembola, 695 – 696
Copepoda, 916 – 917,919 – 920
Decapoda
crayfish, 969 – 970,972 – 973
shrimp, 968 – 969
Ephemeroptera, 660 – 661
Euhirudinea, 478 – 479
Gastropoda, 304 – 305
Gastrotricha, 185 – 186
Hydrachnida, 568
Nematoda, 7,260 – 261
Nematomorpha, 288,290
Odonata, 665
Oligochaeta, 439 – 440
Ostracoda, 813 – 816
Plecoptera, 668
Porifera
dormancy, 107 – 108
growth, 108 – 109
overview, 5,107
Protozoa
encystment, 60
senescence and sex, 60 – 61
sexual maturity, 60
Sphaeriidae, 369 – 370
Trichoptera, 671
Turbellaria, 158 – 159
Unionoidea, 362 – 363
Life table, cladocera population regulation,
863
Limnephilidae, see Trichoptera
Listomatea, anatomy, 55
Locomotion
Bivalvia, 335 – 336
Ostracoda, 823
Rotifera, 196,204 – 205
Turbellaria, 157 – 158
Lophophore, bryzoans, 505 – 506,509
Lotic environments
basin morphometry, 20
classification of streams, 20
current velocity and effects on invertebrates, 23
ecosystem changes along stream course
feeding groups and food resources,
27 – 28
large rivers, 29 – 30
patterns along continuum, 27 – 28
river continuum concept, 27 – 28,30
source of river effects, 28 – 29
insect communities
drift of insects, 741 – 742
flow effects, 737 – 738
functional feeding groups, 738 – 740
recolonization mechanisms, 738 – 739
river continuum concept, 740 – 741
substrates, 736 – 737
ionic content, 26 – 27
oxygen concentrations, 19,26
permanence, 19 – 20
pH effects, 27
springs as source, 28 – 29
stream discharge calculation, 20,22
substrate types, 23 – 25
temperature fluctuations and effects on
invertebrates, 19,25 – 26
Lotka – Volterra equations, 61 – 62
Lumbriculidae, see Oligochaeta
Lutrochidae, see Coleoptera
Lymnaeidae, see Gastropoda
Macromiidae, see Odonata
Macrophyte, grazing by insects, 752
Macrothricoidea, see Cladocera
Mastax, Rotifera, 201
Mating, see Reproduction
Maxillae
cladocera, 850
Crustacea, 776
Ostracoda, 812
Maxillules, Crustacea, 776
Mayfly, see Ephemeroptera
Megaloptera, see also Insect, aquatic
Corydalidae, 677
literature references to species keys,
677
overview, 676 – 677
Sialidae, 677
taxonomic key to larvae, 707
Megasclere, sponge spicule, 102 – 103
Mermithidae, see Nematoda
Mesoveliidae, see Heteroptera
Metretopodidae, see Ephemeroptera
Micromelaniidae, see Gastropoda
Microsclere, sponge spicule, 103,117
Midge, see Diptera
Moinidae, see Cladocera
Molannidae, see Trichoptera
Mollusca, see also Bivalvia; Gastropoda
classes, 8 – 9
Molting
cladocera, 854
Crustacea, 780
Decapoda, 957 – 958
Nematoda, 261
Ostracoda, 815
Monogononta, see Rotifera
Mouth, Bivalvia, 341
Mucocyst, Protozoa, 51
Mucus sheath, Rotifera, 227 – 228
Muscidae, see Diptera
Musculature
Nematoda, 258
Rotifera, 202 – 203
Tardigrada, 535
Myxophaga, see Coleoptera
Naididae, see Oligochaeta
Naucoridae, see Heteroptera
Nematocera, see Diptera
Nematocysts, Cnidaria, 137 – 138,140
Nematoda
anatomy
cuticle, 256,258
excretory system, 259
hypodermis, 258
intestine, 258 – 259
morphology, 256
musculature, 258
nervous system, 259
overview, 8,256
pharynx, 258
reproductive organs, 260
sensory organs, 259 – 260
stoma, 258
collection
extraction, 265 – 266,268
sampling, 265
culture, 269
development, 260
feeding, 261 – 263
fossil record, 261 – 262
functional classification, 255
habitat, 262 – 265
hatching, 260 – 261
life cycle, 7,260 – 261
molting, 261
predation, 265
resistant stages, 261
respiration, 259
Subject Index
specimen preparation for identification,
268 – 269
systematic arrangement of genera, 295
taxonomic key
Mermithidae, 278,280
non-Mermithidae, 270,274,276,278
toxicology, 265
Nematomorpha
anatomy
cuticle, 282 – 283
digestive tract, 283
overview, 8,282
reproductive organs, 283
collection, 290 – 291
evolution, 280 – 282
fossil record, 280
larva, 284
life cycle, 288,290
oviposition, 283
parasitism, 282,288,290
specimen preparation for identification,
291
taxonomic key, 291 – 292
Nemertea
anatomy, 6 – 7,173
behavioral ecology, 175
classification, 174
collection, 176
culture, 176
distribution, 175
ecosystem role, 175 – 176
feeding, 174
habitat, 7,175
physiological adaptations, 175
preservation of specimens, 176
prospects for research, 176
reproduction, 174 – 175
taxonomic key, 176
Nemouridae, see Plecoptera
Neoephemeridae, see Ephemeroptera
Nepidae, see Heteroptera
Nepomorpha, see Heteroptera
Neritinidae, see Gastropoda
Nervous system
Bivalvia, 343 – 344
Branchiobdellida, 457
bryzoans, 506 – 507
Copepoda, 916
Crustacea, 779
Decapoda, 954 – 955
Gastrotricha brain, 182
Hydrachnida, 567 – 568
Nematoda, 259
Ostracoda, 812
Rotifera, 203 – 204
Tardigrada, 535
Neuroptera, see also Insect, aquatic
diversity, 681
Sisyridae, 681
Nitrogen excretion
Bivalvia, 340,351
Copepoda, 914
Crustacea, 777
Noteridae, see Coleoptera
Notonectidae, see Heteroptera
Notostraca, see Branchiopoda
Nymphomyiidae, see Diptera
Odonata, see also Insect, aquatic
Anisoptera (dragonfly)
Aeshinidae, 666
Cordulegastridae, 666
Corduliidae, 666 – 667
Gomphidae, 667
Libellulidae, 667
Macromiidae, 667
Petaluridae, 667
ecological impact, 665
larva, 665 – 666
life cycle, 665
literature references to species keys, 666
reproduction, 665
taxonomic key to larvae, 700
Zygoptera (damselfly)
Calopterygidae, 667
Coenagrionidae, 667
Lestidae, 667
Protoneuridae, 667 – 668
Odontoceridae, see Trichoptera
Oligochaeta, see also Annelida
anatomy
chaetae, 435 – 436,443
clitellum, 432,434
eye spots, 437 – 438
gills, 436
morphology, 432
pharyngeal glands, 437
prostomium, 434 – 435
reproductive organs, 432 – 434
sensory organs, 436 – 437
collection, 444
culture, 444 – 445
defecation rates, 440 – 441
desiccation tolerance, 438
distribution, 438 – 439
diversity, 438 – 439
evolution
clitellate taxa, 443 – 444
Enchytraeidae, 441
Haplotaxidae, 441
Lumbriculidae, 443
Naididae, 441,443
phylogenetic tree of Annelida, 442
Tubificidae, 441,443
feeding, 437,440
habitats, 10,431 – 432,438
life cycle, 439 – 440
reproduction, 434,439 – 440
respiration, 437
salt tolerance, 437
specimen preparation for identification,
445 – 446
taxonomic keys
families, 446 – 448
Lumbriculidae, 448 – 450
Naididae, 450 – 452
Tubificidae, 452 – 454
Oligohymenophorea, anatomy, 55
Oligoneuriidae, see Ephemeroptera
1031
Onychopoda, see Cladocera
Oscula, sponges, 101
Osmoregulation
Bivalvia
carbonic anhydrase role, 353
hemolymph osmolarity, 353
hormonal control, 354
ion transport, 352 – 353
salt tolerance, 352
Crustacea, 777,779
Decapoda, 959
Osphradia, bivalves, 344
Ostracoda, see also Crustacea
anatomy
body, 809
circulatory system, 812
digestive system, 812
eye, 812
head, 809,811
legs, 812
maxillae, 812
nervous system, 812
reproductive organs, 812 – 813
shell morphology
external, 808 – 809
internal, 809
applications
environmental tolerance, 827
paleolimnology, 827
trace element analysis in shells,
827 – 828
collection, 825 – 826
coloration, 823
controversies in taxonomy, 828 – 829
culture, 826 – 827
distribution
benthic nonswimmers, 816
benthic swimmers, 816
egg, 813,822
embryology, 814
feeding, 823 – 824
fossils, 807 – 808,827
habitats
bromeliads, 821
carbonate content, 816,819
chemical water types, 817
diversity, 816
lakes, 821
oxygen concentrations, 818 – 819
pH tolerance, 818
population regulation, 824 – 825
salinity, 817 – 818
sediment – water interface, 822
solute composition, 818
streams, 821
temperature, 819 – 821
underground waters, 821 – 822
hatching and seasonality, 814 – 815
intraspecific competition, 816
life cycle, 813 – 816
locomotion, 823
molting, 815
overview, 13
parasitism, 825
Podocopida, 829
1032
Subject Index
Ostracoda (continued)
predation, 824
prospects for research, 828
reproduction, 813 – 814
resting stage and torpidity, 822 – 823
specimen preparation, 826
taxonomic keys
Cypridoidea, 829 – 830,832 – 835
Cytheroidea, 835 – 838
toxicology, 825
ultraviolet-B tolerance, 825
Oxygen concentration
aquatic insect effects, 731 – 732
Bivalvia effects, 349 – 350,384
Copepoda adaptation, 921 – 922,927
Euhirudinea
hyperoxia tolerance, 473
hypoxia tolerance, 472 – 473
Gastropoda hypoxia adaptation, 302
noncladoceran adaptation, 891
Protozoa effects, 56 – 57
Rotifera tolerance, 206
Oxygen consumption, bivalves
biomass relationship, 346
growth and reproduction relationship, 347
hypoxia response, 348 – 349
pollutant effects, 348
seasonal cycles, 344 – 346
Palaemonidae, see Decapoda
Paleolimnology
cladocera applications, 864 – 865
Ostracoda applications, 827
Porifera, 114
Papillae
Acanthobdellidae, 468
Euhirudinea, 468
Parasitism
Acanthobdellidae, 466 – 467,474
Bivalvia parasites, 384
Branchiobdellida parasitism and parasites, 458 – 459
bryzoan parasites, 513
Copepoda, 929
Decapoda, 974
Gastropoda, 308 – 309
Hydrachnida parasitism of insects by
larva
attachment and engorgement, 574 – 575
detachment, 575
ecological impact
host populations, 580 – 581
individual hosts, 580
intraspecific effects on same host,
581
emergence of larvae, 572 – 573
host selection, 569 – 570
infection, 574
recognition of host, 573
seeking and locating hosts, 573
site selection, 570 – 572
success rates, 569
Nematomorpha, 282,288,290
Ostracoda, 825
Rotifera, 228 – 229
Turbellaria, 161
Unionoidea glochidium on fish,
363 – 364,368
Parisitiformes, features, 649
Peltoperlidae, see Plecoptera
Peracarida, see also Crustacea
Amphipoda, 784
evolution, 785 – 786
feeding, 788 – 790
Isopoda, 784 – 785
orders, 784
overview, 12 – 13
predation, 789 – 790
reproduction, 791
taxonomic key, 795 – 796
Perlidae, see Plecoptera
Perlodidae, see Plecoptera
Petaluridae, see Odonata
pH
aquatic insect effects, 733
Bivalvia
distribution effects, 360
population effects, 383 – 384
cladocera species diversity effects, 857
crayfish effects, 977 – 978
Crustacea tolerance, 781
Gastropoda effects, 309
Rotifera tolerance, 206
Phagocytosis, Porifera, 102
Pharyngeal glands, Oligochaeta, 437
Pharynx, Nematoda, 258
Phenemics, taxonomic classification, 1
Philopotamidae, see Trichoptera
Phryganeidae, see Trichoptera
Phyllopharyngea, anatomy, 55
Phylogeny, see Evolution
Physidae, see Gastropoda
Piscicolidae, see Euhirudinea
Pisciforma, see Ephemeroptera
Planorbidae, see Gastropoda
Plecoptera, see also Insect, aquatic
biomonitoring studies, 763 – 764
Euholognatha
Capniidae, 669
Nemouridae, 669 – 670
Taeniopterygidae, 670
feeding, 668 – 669
larva, 668
life cycle, 668
literature references to species keys, 669
Systellognatha
Chloroperlidae, 670
Peltoperlidae, 670
Perlidae, 670
Perlodidae, 670
Pteronarcyidae, 670
taxonomic key to larvae, 700 – 703
Pleidae, see Heteroptera
Pleuroceridae, see Gastropoda
Podocopida, see Ostracoda
Poecilostomatoida, see Copepoda
Polycentropodidae, see Trichoptera
Polymitarcyidae, see Ephemeroptera
Polyphaga, see Coleoptera
Pomatiopsidae, see Gastropoda
Ponds, see Lentic ecosystems
Porifera
anatomy
digestion, 102
external morphology, 98,100
overview, 5,97 – 98
skeleton
collagen, 102,104
siliceous spicules, 102 – 104
water processing system, 100 – 102
biochemistry, 105
classification by spicules, 117 – 127
collection, 115
culture, 115
distribution, 106 – 107,117 – 127
diversity, 105 – 106
ecology
algal symbionts, 110 – 111
competition for substrates, 112 – 113
dispersal among habitats, 109
feeding, 110 – 112
functional role in ecosystems, 114
infaunal organisms, 113 – 114
paleolimnology, 114
predation of sponges, 113
silica effects on skeleton structure, 112
substrate availability, 109 – 110
evolutionary relationships, 114 – 115
life cycle
dormancy, 107 – 108
growth, 108 – 109
overview, 5,107
phagocytosis, 102
reproduction
asexual, 104 – 105,108 – 109
sexual, 104,108
specimen preparation for identification,
115 – 116
taxonomic key, 127 – 130
Potamanthidae, see Ephemeroptera
Predation
Acanthobdellidae, 480 – 481
aquatic insects
fish predation, 753 – 754
insects as predators, 752 – 754
lentic versus lotic environments,
752 – 754
Bivalvia, 385 – 386
bryzoans, 513
cladocera, 863 – 864
Copepoda, 920,927
crayfish, 978 – 981
Crustacea, 789 – 790
Euhirudinea, 480 – 481
Gastropoda, 297,310 – 312
Gastrotricha, 187
Hydrachnida, 582
Nematoda, 265
Ostracoda, 824
Peracarida, 789 – 790
Porifera, 113
Subject Index
Protozoa, 62,67 – 68
Rotifera, 226 – 228
Turbellaria, 161
Prostomatea, anatomy, 55
Prostomium, Oligochaeta, 434 – 435
Protonephridium, Rotifera, 204
Protoneuridae, see Odonata
Protozoa
anatomy
amoebas, 53 – 54
ciliatea, 54 – 55
flagellates, 51 – 52
Heliozoa, 54
cellularity, 45
collection, 69 – 70
culture, 69 – 70
distribution, 59
diversity, 58 – 59
ecology
algivores, 65,67
bacterivores, 64 – 65
ecosystem role, 4 – 5,43
epizooic associations with animals,
63 – 64
histophagous ciliates, 67
interspecific interactions, 61 – 62
intraspecific interactions, 61
metazoan predators, 67 – 68
nutrient cycling, 68
predatory interactions, 62
symbionts, 62 – 63
environmental physiology
contaminants, 57 – 58
energetics, 58
factors affecting distribution, 55 – 56
oxygen, 56 – 57
temperature, 56
evolutionary relationships, 68 – 69
life cycle
encystment, 60
senescence and sex, 60 – 61
sexual maturity, 60
organelles
cilia, 47 – 48
contractile vacuoles, 46 – 47
extrusomes, 48,51
flagella, 47 – 48
food vacuoles, 45 – 46
phyla, 4,44 – 45
reproduction
fission, 59
generation time, 59 – 60
sexual versus asexual reproduction, 60
taxonomic categories, 44 – 45
taxonomic keys
ciliates, 78 – 79,81 – 83,85,87,89
flagellates, 71
functional groups, 70 – 71
overview, 70
pseudopodia protozoa, 71,73 – 78
Psephenidae, see Coleoptera
Pseudironidae, see Ephemeroptera
Psychodidae, see Diptera
Psychomyiidae, see Trichoptera
Pteronarcyidae, see Plecoptera
Ptilodactylidae, see Coleoptera
Ptychopteridae, see Diptera
Pupa, Diptera, 689
Q10 value, 345 – 346
Radula, Gastropoda, 300
RCC, see River continuum concept
rDNA, see Ribosomal DNA
Rearing, see Culture
Regeneration, Hydra, 143,145
Removal-summation method, secondary
production measurement, 758
Reproduction
Acanthobdellidae, 477 – 479
Bivalvia
egg, 343
sperm, 343
temperature regulation, 343
Branchiobdellida, 458
Branchiura, 784
bryzoans
asexual budding, 509 – 510
fission, 510
gametogenesis, 513
sexual, 509
cladocera, 852 – 854,859 – 860
Cnidaria, 138 – 140
Copepoda, 916 – 917,919
Decapoda
crayfish, 969 – 970,972 – 973
gonads, 958
pheromones, 958,983
shrimp, 968 – 969
Ephemeroptera, 661
Euhirudinea, 471,477 – 479
Gastropoda, 299 – 300,304 – 305
Gastrotricha, 185 – 186
Hydrachnida
pheromones, 577
sex ratios, 578
spermatophore transfer
association during transfer, 576 – 577
copulation during transfer, 577 – 578
dissociation during transfer, 576
Nematomorpha oviposition, 283
Nemertea, 174 – 175
Odonata, 665
Oligochaeta, 434,439 – 440
Ostracoda, 813 – 814
Peracarida, 791
Porifera
asexual, 104 – 105,108 – 109
sexual, 104,108
Protozoa
fission, 59
generation time, 59 – 60
sexual versus asexual reproduction, 60
Rotifera
cyclical parthenogenesis, 214 – 215
mating behavior, 217
1033
resting eggs, 215 – 217
Sphaeriidae, 368 – 369
Tardigrada
modes, 536 – 537
apparatus, 537
mating and fertilization, 537
Reproductive organs
Acanthobdellidae, 470
Bivalvia, 341 – 343
Branchiobdellida, 457
cladocera, 852
Copepoda, 914,916
Crustacea, 779 – 780
Euhirudinea, 470
Gastropoda, 299 – 300
Gastrotricha, 182
Hydrachnida, 568
Nematoda, 260
Nematomorpha, 283
Oligochaeta, 432 – 434
Ostracoda, 812 – 813
Rotifera, 204
Tardigrada, 537
Respiration
Acanthobdellidae, 472
Coleoptera, 682 – 683
Decapoda, 955 – 957
Euhirudinea, 472
Nematoda, 259
Oligochaeta, 437
Respiratory system
Crustacea, 777
Gastropoda, 301
Hydrachnida, 567
Retrachaeta, see Ephemeroptera
Rhabdoids, Turbellaria, 157,162
Rhyacophilidae, see Trichoptera
Ribosomal DNA (rDNA), Gastropoda phylogenetic analysis, 313 – 314
River continuum concept (RCC), 740 – 741
Rivers, see Lotic environments
Rossianidae, see Trichoptera
Rotifera
aging and senescence, 217 – 218
anatomy
corona, 198 – 199,201
distinguishing features, 195
foot, 197
mastax, 201
morphology, 197 – 198
muscular system, 202 – 203
nervous system, 203 – 204
protonephridium, 204
reproductive organs, 204
spines, 226 – 228
trophi, 201 – 202,234
anhydrobiosis and rehydration, 207
biogeography, 210 – 211
classes, 197,233
clearance rates, 224 – 225
collection, 230 – 231
colonization, 196,211 – 213
competition with other zooplankton,
225 – 226
1034
Subject Index
Rotifera (continued)
culture, 231
distribution and density, 195 – 196,208
ecology overview, 7
ecosystem role, 224 – 225
feeding behavior, 223 – 224
fish aquaculture application, 229 – 230
locomotion, 196,204 – 205
mucus sheaths, 227 – 228
parasitism, 228 – 229
phenotypic variation and mechanisms,
207 – 208
polyploidy, 205
population dynamics
dynamics, 221 – 222
genetic variation, 222 – 223
life table analysis, 218 – 221
population movements, 208 – 209
predator – prey interactions, 226 – 228
reproduction
cyclical parthenogenesis, 214 – 215
mating behavior, 217
resting eggs, 215 – 217
sessile species, 213 – 214
specimen preparation for identification,
231 – 233
systematics
Bdelloidea, 233
Monogononta, 233 – 234
Seisonidea, 233
taxonomic key,
234 – 237,239 – 245,247 – 248
tolerance
oxygen deprivation, 206
pH, 206
salt, 206
temperature, 205 – 206
toxicology, 206 – 207
Roundworm, see Nematoda
Salinity
aquatic insect effects, 733 – 734
cladocera species diversity effects,
857
Crustacea tolerance, 780 – 781
Euhirudinea tolerance, 473 – 474
noncladoceran adaptation, 891
Oligochaeta tolerance, 437
Rotifera tolerance, 206
Salt tolerance, see Salinity
Schizopyrenida, anatomy, 53 – 54
Sciomyzidae, see Diptera
Scirtidae, see Coleoptera
Seisonidea, see Rotifera
Sensory organs
Bivalvia, 344
Copepoda, 916
Crustacea, 781 – 783
Decapoda, 955
Nematoda, 259 – 260
Oligochaeta, 436 – 437
Turbellaria, 157
Sericostomatidae, see Trichoptera
Setisura, see Ephemeroptera
Shell
Bivalvia synthesis, 331,334 – 335
Gastropoda, 298 – 299
Ostracoda morphology
external, 808 – 809
internal, 809
Unionoidea growth, 366
Shrimp, see Decapoda
Sialidae, see Megaloptera
Sididae, see Cladocera
Simuliidae, see Diptera
Siphlonuridae, see Ephemeroptera
Sisyridae, see Neuroptera
Size frequency method, secondary production measurement, 758
Skeleton
Crustacea, 776
Porifera
collagen, 102,104
siliceous spicules, 102 – 104
Snail, see Gastropoda
Specimen preparation
aquatic insects, 660
Bivalvia, 394 – 395
Branchiobdellida, 459
bryzoans, 516 – 517
Collembola, 696
Copepoda, 930 – 931
Crustacea, 795
Decapoda, 986 – 987
Euhirudinea, 486 – 487
Gastropoda, 315 – 316
Hydrachnida, 585
Nematoda, 268 – 269
Nematomorpha, 291
Nemertea, 176
noncladocerans, 895
Oligochaeta, 445 – 446
Ostracoda, 826
Porifera, 115 – 116
Rotifera, 231 – 233
Tardigrada, 541
Turbellaria, 155,163
Sphaeriidae, see also Bivalvia
evolution, 390,392 – 393
growth rates, 370
habitat and life history, 369 – 370
life cycle, 368
reproduction, 368 – 369
Spicipalpia, see Trichoptera
Spinicaudata, see Branchiopoda
Sponges, see Porifera
Spongillafly, see Neuroptera
Springtail, see Collembola
Statoblast, bryzoans, 506,510,517
Statocyst
bivalves, 344
Decapoda, 955
Stoma, Nematoda, 258
Stomach, Bivalvia, 341
Stonefly, see Plecoptera
Stratiomyidae, see Diptera
Streams, see Lotic environments
Suckers
Acanthobdellidae, 468 – 469
Euhirudinea, 468 – 469,475
Swamps, see Lentic ecosystems
Syrphidae, see Diptera
Systellognatha, see Plecoptera
Systematic zoology, taxonomic classification, 2
Tabanidae, see Diptera
Taeniopterygidae, see Plecoptera
Tanyderidae, see Diptera
Tardigrada
anatomy
claws, 530 – 532
cuticle, 528 – 530
digestive system
buccal aparatus, 532 – 534
esophagus, 534
hindgut, 534
midgut, 534
overview, 532
excretory system, 535
morphology, 527 – 528
musculature, 535
nervous system, 535
reproductive organs, 537
collection, 541
cryptobiosis
encystment, 535 – 536
anoxybiosis, 536
cryobiosis, 536
osmobiosis, 536
anhydrobiosis, 536
dissemination, 540
distribution, 540
evolution, 527
habitats, 537 – 539
historical perspective of study, 527
overview, 11
population density, 539 – 540
reproduction
apparatus, 537
mating and fertilization, 537
modes, 536 – 537
specimen preparation and microscopy,
541
taxonomic key, 542 – 544
Taxonomic keys
Acanthobdellidae, 489
Bivalvia
Corbiculacea, 398 – 400
superfamilies, 397
Unionoidea, 400 – 403,408 – 415
Branchiobdellida, 459 – 460
Branchiopoda
cladocera
Bosminidae, 881
Chydoridae, 874 – 878
Daphniidae, 879 – 881
families, 871,873
Halopediidae, 873
Haplopoda, 885,887
Macrothricoidea, 883
Moinidae, 879 – 881
Onychopoda, 885,887
Subject Index
Sididae, 873 – 874
noncladocera, 895,897,900
bryzoans, 517 – 523
Cnidaria, 152 – 153
Collembola, 716 – 717
Copepoda
Calanoida, 934 – 935
Cyclopoida, 936 – 937
Harpacticoida, 937 – 938
Crustacea orders, 795 – 796
Decapoda, 987 – 989
Euhirudinea, 489 – 497
Gastropoda, 316,318,320,323
Gastrotricha, 189 – 191
Hydrachnida
adults, 638 – 648
larvae, 633 – 638
overview, 585 – 586
insects, aquatic
Coleoptera
adults, 708,710 – 711
larvae, 712 – 714
Diptera larvae, 714 – 716
Ephemeroptera larvae, 697 – 700
Heteroptera, 708
Megaloptera larvae, 707
Odonata larvae, 700
orders, 696 – 697
Plecoptera larvae, 700 – 703
Trichoptera larvae, 704 – 705,707
major taxa of freshwater invertebrates,
16 – 18
Nematoda
Mermithidae, 278,280
non-Mermithidae, 270,274,276,278
Nematomorpha, 291 – 292
Nemertea, 176
Oligochaeta
families, 446 – 448
Lumbriculidae, 448 – 450
Naididae, 450 – 452
Tubificidae, 452 – 454
Ostracoda
Cypridoidea, 829 – 830,832 – 835
Cytheroidea, 835 – 838
Porifera, 127 – 130
Protozoa
ciliates, 78 – 79,81 – 83,85,87,89
flagellates, 71
functional groups, 70 – 71
overview, 70
pseudopodia protozoa, 71,73 – 78
Rotifera, 234 – 237,239 – 245,247 – 248
Tardigrada, 542 – 544
Temperature
aquatic insect effects, 732 – 733
Bivalvia
distribution effects, 361
population effects, 384
Branchiopoda
cladocera metabolic rate effects, 855
noncladoceran adaptation, 891
Copepoda
adaptation, 921
population regulation effects, 926 – 927
Crustacea tolerance, 781
Decapoda adaptation, 958
Gastropoda effects, 301 – 302
Protozoa effects, 56
Rotifera tolerance, 205 – 206
Testacealobosia, anatomy, 53
Thaumaleidae, see Diptera
Thiaridae, see Gastropoda
Thorax, Copepoda, 912
Tipulidae, see Diptera
Toxicology
cladocera, 867,869
Decapoda, 959 – 960
Euhirudinea, 485 – 486
Hydrachnida, 582 – 583
Nematoda, 265
noncladocerans, 894 – 895
Ostracoda, 825
Rotifera, 206 – 207
Trichoptera, see also Insect, aquatic
Annulipalpia
Dipseudopsidae, 672 – 673
Ecnomidae, 673
Hydropsychidae, 673
Philopotamidae, 673
Polycentropodidae, 673
Psychomyiidae, 673
Xiphocentronidae, 673
biomonitoring studies, 763 – 764
diversity, 670
ecological impact, 672
identification, 671
Integripalpia
Apataniidae, 674
Beraeidae, 674
Brachycentridae, 674
Calamoceratidae, 674
Goeridae, 674
Helicopsychidae, 674
Lepidostomatidae, 674
Leptoceridae, 674 – 675
Limnephilidae, 675
Molannidae, 675
Odontoceridae, 675
Phryganeidae, 675
Rossianidae, 675
Sericostomatidae, 675
Uenoidae, 675 – 676
larva
adaptations, 671
collection, 671 – 672
pupate, 671
life cycle, 671
literature references to species keys,
672
Spicipalpia
Glossosomatidae, 676
Hydrobiosidae, 676
Hydroptilidae, 676
Rhyacophilidae, 676
taxonomic key to larvae, 704 – 705,
707
Trinomen, taxonomic classification, 2
Trophi, Rotifera, 201 – 202,234
Tubificidae, see Oligochaeta
1035
Turbellaria
anatomy
ciliated epidermis, 157
eyes, 157
morphology, 156 – 157
overview, 6 – 7
rhabdoids, 157,162
sensory organs, 157
behavioral ecology, 160
collection, 163
community ecology, 162
culture, 163
distribution, 156,159 – 160,162
ecosystem role, 161 – 162
endosymbiosis, 162
environmental physiology, 158
feeding, 160 – 161
habitat, 7
life history, 158 – 159
locomotion, 157 – 158
population regulation and density, 161
predation and parasitism, 161
preservation of specimens, 155,163
species list, 179 – 180
status in Animal kingdom, 156
Uenoidae, see Trichoptera
Underground aquatic habitats
caves
food sources, 31
karst topography in United States, 31
life cycle of species, 33
protection, 32 – 33
species distribution, 31 – 32
hyporheic zone, 30 – 31
phreatic zone, 31
Unionoidea, see also Bivalvia
conservation biology, 368
distribution, 402
ecosystem role, 387
evolution, 390 – 393
gill calcium phosphate concretions,
350,352
glochidium
collection, 394
fecuindity, 364,366
fish parisitism, 363 – 364,368
forms, 363
host food mimicry, 364
host reduction effects, 385
size versus number released, 366
growth rates, 366 – 367
life cycle, 362 – 363
life span, 366 – 367
shell growth, 366
standing crop biomass rate versus biomass production, 367
taxonomic key, 400 – 403,408 – 415
Valvatidae, see Gastropoda
Vascular system
Acanthobdellidae, 471 – 472
Branchiobdellida, 457
Euhirudinea, 471 – 472
1036
Subject Index
Veliidae, see Heteroptera
Visceral mass, Gastropoda, 299
Vitamin E, Rotifera effects
longevity, 218
mictic reproduction induction, 215
Viviparidae, see Gastropoda
Water beetle, see Coleoptera
Water flea, see Cladocera
Water mite, see Hydrachnida
Web sites, freshwater invertebrate taxonomy, 3 – 4
Wetlands, see Lentic ecosystems
Xiphocentronidae, see Trichoptera
Zebra mussel, see Dreissena polymorpha
Zooid, bryzoans, 505 – 506
Zoological ranks, 3
Zygoptera, see Odonata
Taxonomic Index
Abedus, 680
Ablabesmyia janta, 384
Abramis brama, 923
Abrochta, 236
Acalyptonotus, 552, 559, 579,
635, 647
neoviolaceus, 602, 628
Acanthamoeba, 73, 74
Acanthobdella, 442, 444
peledina, 17, 466, 467, 470, 471, 472,
479, 486, 489
Acanthocyclops, 921, 937
brevispinosus, 949
capillatus, 949
carolinianus, 949
columbiensis, 949
exilis, 949
montana, 949
parasensitivus, 949
parvulus, 949
pennaki, 949
robustus, 928, 949
venustoides, 949
vernalis, 949
Acanthocystis, 78
turfacea, 77
Acanthodiaptomus, 929, 935
denticornis, 948
Acantholeberis, 883
curvirostris, 884
Acartia, 923
sinensis, 934
Acella haldemani, 320, 321
Acentrella, 662
Acerpenna, 662
Acherontacarus, 553, 563, 569, 606, 633,
638
Achromadora, 265, 276
Acilus, 683
Acineta, 81
limnetis, 79
Acipenser
fulvesens, 385
guldenstadti, 149
ruthenus, 148, 149
Acroloxus, 303, 316, 317
Acroneuria, 658, 730
Acroperus, 874, 876
harpae, 875
Actinarctus, 538
Actinobdella, 488, 490
annectens, 490
inequiannulata, 474, 488, 490
Actinobolina, 51, 85
radians, 84
Actinolaimus, 274
Actinomonas, 48
Actinonaias, 412
Actinophrys, 78
sol, 61, 77
Actinosphaerium, 54, 62, 78
eichhorni, 77
Acutogordius, 292
Acyclus, 235
Adineta, 228, 235, 236
Adorybiotus, 529, 531, 532, 543
1037
1038
Taxonomic Index
Aedes, 730
cinereus, 574
communis, 572
excrucians, 574
punctor, 572
Aeolosoma, 447
Aeromonas hydrophila, 470
Aeshna, 730
eremita, 782
Agabus, 683
Agarodes, 675
Aglaodiaptomus, 935
atomicus, 948
clavipes, 948
clavpoides, 948
conipedatus, 948
dilobatus, 948
forbesi, 948
kingsburyae, 948
leptopus, 948
lintoni, 948
marshianus, 948
pseudosanguineus, 948
saskatchewanensis, 948
spatulocrenatus, 948
stagnalis, 948
Agneta, 669
Agostocaris, 961
Agriodrilus, 443
Alaimus, 278
Alasmidonta, 402, 403, 409, 411, 413, 414
mccordi, 357
robusta, 357
undulata, 404
wrightiana, 357
Alathyria jacksoni, 348
Albaxona, 558, 646
nearctica, 622
Albertathyas, 553, 640
montana, 609
Albia, 557, 622, 638, 647
neogaea, 598
Alboglossiphonia, 488
heteroclita, 492, 494
Allocapnia, 703
Allodero, 439, 451
Allonais, 438, 451, 452
Alloperla, 702
Alona, 847, 852, 874, 877
acutirostris, 878
affinis, 878
bicolor, 849, 877
intermedia, 878
verrucosa, 878
Alonella, 874, 878
dadayi, 878
excisa, 878
exigua, 862, 878
hamulata, 878
nana, 878
pulchella, 878, 879
Alonopsis, 874, 876
elongata, 862, 875
Alosa sapidissma, 385
Amblema, 402, 413
plicata, 361, 381, 404, 409
Ambrysus, 678
Ambystoma
barbouri, 791
maculosum, 479, 480
tigrinum, 479
Ameletus, 699
Amerothyasella, 553, 640
appalachiana, 611
Ametropus, 699
Amnicola, 307, 309, 320
limosa, 304
Amoeba, 5, 43, 53, 54, 65, 74
proteus, 53, 74
Amoenacarus, 559, 645
dixiensis, 628
minimus, 628
Amphibolella, 169
spinulosa, 179
Amphibolus, 529, 531, 532, 533, 539, 543
Amphichaeta, 436, 450
Amphidelus, 278
Amphileptus, 87
claparedi, 86
Amphinemura, 703
Amphisiella, 83
oblonga, 82
Amphizoa, 711, 713
Anabaena
affinis, 225
oscillariodes, 395
Anacaena, 684
Anacanthoderma, 191
Anas platyrhynchos, 892
Anatonchus, 278
Anatopynia, 161
Anax, 701
junius, 666
Anchistropus, 140, 862, 874, 878
minor, 879
Anchytarsus, 684, 713
Ancylus fluviatilis, 306, 310
Anheteromeyenia, 130
argyrosperma, 116, 117, 130
ryderi, 106, 117, 118, 130
Ankistrodesmus, 395, 396, 870
Ankylocythere, 830, 836
Ankyrodrilus, 460
Anodonta, 363, 402, 403, 409, 412
anatina, 348, 363, 367, 376, 384, 391
cygnea, 335, 339, 340, 341, 353, 354,
382, 391
grandis, 345
imbecelis, 396
implicata, 354
woodiana, 353, 366
Anodontoides, 402, 404, 412
ferrussacianus, 409
Anonchus, 276
Anopheles, 691, 715, 730
crucians, 573
Anthopotamus, 698
Antiprodrilus, 443
Antrobia, 320
Anuraeopsis, 210
fissa, 205, 217
Apartronemertes, 174
Apatania, 672
Aphanolaimus, 276
Aphaostracon, 320
Aphelenchoides, 263, 270, 273
Aphelenchus, 263, 270
Apheviderulix, 553, 639
santana, 607
Aplexa, 303, 304, 318
elongata, 302, 306, 319
Aplodinotus grunniens, 385
Apocyclops, 937
dimorphus, 949
panamensis, 949
spartinus, 949
Apodibius, 529, 532, 542
Apolodinotus grunniens, 474
Apsilops, 730
Aqabus, 711, 713
Aqarodes, 706
Aquarius, 678
Aranimermis, 278, 280
Arcella, 67, 74
vulgaris, 75
Archilestes, 667
Arcidens, 402
confragosa, 404, 409, 410, 413
Arcteonais, 438
lomondi, 450
Arctodiaptomus, 935
arapahoensis, 948
bacillifer, 948
dorsalis, 913, 916, 948
floridanus, 948
kurilensis, 948
saltillinus, 948
Arctodrilus wulikensis, 438, 454
Arenohydracarus, 559, 579, 646
Arenotus, 185
Argia, 730
Argulus, 13, 783, 784, 787, 795
maculosus, 784
Arizonacarus, 556, 643
chircahuensis, 616
Arkansia wheeleri, 404, 409
Armiger, 318
crista, 317
Arrenurus, 552, 559, 567 – 569,
572 – 576, 579, 580, 628, 631,
635, 644
angustilimbatus, 572
cuspidator, 575
danbyensis, 573, 580
delawarensis, 573
fissicornis, 629
kenki, 572, 574
manubriator, 578, 582
planus, 575, 578, 604
pseudocylindratus, 629
pseudosuperior, 569, 580
ventropetiolatus, 575
Taxonomic Index
Artemia, 150, 229, 889 – 895, 897, 931
franchiscana, 888, 891
salina, 38, 891
Artemiopsis, 889, 892, 893, 897
stephanssoni, 896
Arthroplea, 699
Asajirella, 510
Ascetocythere, 830, 836
Ascomorpha, 202, 204, 247, 248
Ascomorphella volvocicola, 244, 246
Ascophora, 170
elegantissima, 180
Asellus, 163, 785, 786, 794, 796
aquaticus, 785
Askensia, 85
volvox, 84
Aspidiophorous, 189, 190
Aspidisca, 81
costata, 80
Asplanchna, 198, 201 – 205, 215, 219,
223 – 228, 232, 239, 240
brightwelli, 205, 208, 217, 218, 219,
222, 223
girodi, 205, 216, 217, 223, 227
intermedia, 208
priodonta, 209, 222
sieboldi, 204, 208
Asplanchnopus, 201, 227, 239
calyciflorus, 203
Astacus, 967
Astatumen, 529, 531, 534
Astylozoon, 81
faurei, 80
Atherix, 693, 715
Athernia anthonyi, 314
Atopsyche, 706
Atractides, 556, 574, 579, 618, 637, 643,
647
grouti, 593
Atrochus, 235
Attenella, 662
Attheyella, 921, 926, 938
americana, 950
carolinensis, 950
dentata, 950
dogieli, 950
idahoensis, 950
illinoisensis, 950
nordenskioldi, 950
obatogamensis, 950
pilosa, 950
spinipes, 913, 950
trispinosa, 950
ussuriensis, 950
Aturus, 552, 557, 578, 579, 600, 624, 631,
637, 646
Atylenchus, 270
Atylotus, 690
Aulodrilus, 438, 443, 454, 455
pluriseta, 452
Aulolaimoides, 274
Aulophorus, 439, 451, 451
Australocypris insularis, 823
Austropotamobius, 967
Automate, 961
Axonopsella, 558, 646
bakeri, 621
Axonopsis, 558, 566, 572, 579, 637, 647
beltista, 623
setoniensis, 600
Baetis, 13, 662, 699, 730, 737, 739, 741,
742, 743, 754, 760
Baetisca, 698
Baetodes, 662
Balanonema, 87
biceps, 86
Bandakia, 555, 636, 642
phreatica, 590, 613
Bandakiopsis, 555, 634, 642
fonticola, 589, 613
Barbicambarus, 963, 964, 966, 967, 988
Barbidrilus, 447
Barbouria, 961
Barentsia, 515
Bastiania, 276
Batillipes, 538
Batracobdella, 468, 469, 488
cryptobranchii, 488, 493
michiganensis, 488
paludosa, 488, 492
Bdellergatis, 489
plumbeus, 482, 496
Bdellodrilus, 460
illuminatus, 460
Beatogordius, 292
Belostoma, 680, 709, 730
Beraea, 706
Berosus, 684
Bezzia, 580
Bharatalbia, 557, 624, 646
cooki, 624
surensis, 624
Biapertura, 878
Bicoeca, 71
lacustris, 72
Binucleospora elongata, 825
Biomphalaria, 313
glabrata, 306, 319
Biomyxa, 77
vagans, 76
Birgea, 240
enantia, 237
Birgella, 320
Bithynia tentaculata, 306, 322, 323
Blepharicera, 690, 715
Blepharisma, 43, 61, 62, 83
lateritium, 84
Bodo, 52, 71
caudatus, 72
Boreobdella verrucata, 494
Bosmina, 847, 850, 851, 856, 857, 859,
861, 862, 864, 865, 881
freyi, 2
longirostris, 2, 225, 881, 882
Bosminopsis, 881
deitersi, 882
1039
Bothrioneurum, 434, 435, 439, 455
vejdovskyanum, 454
Bothrioplana, 157, 164
semperi, 159, 169, 179
Bothromesostoma, 161, 171, 173
personatum, 180
Bouchardina, 963, 964, 965, 966, 968, 989
robinsoni, 987
Brachionus, 210, 215, 217, 219, 223, 224,
227, 228, 243, 244
angularis, 223, 232
bidentata, 227
calyciforus, 205, 207, 208, 216, 218,
219, 220, 222 – 227, 229, 232
caudatus, 208
patulus, 205, 217, 232
plicalitis, 39, 196, 197, 204 – 207,
215 – 219, 223, 229, 232, 243
quadridentatus, 223, 223
rubens, 207, 225, 229, 232
urceolaris, 219, 220, 223, 227
Brachycentrus, 672, 706, 738, 743
Brachycercus, 662
Brachypoda, 558, 577, 637, 646
cornipes, 599
oakcreeksensis, 622, 623
setosicauda, 599, 600
Bradleystrandesia, 828
Bradyscela, 235
Branchinecta, 889, 890, 891, 892, 893, 897
campestris, 891
coloradensis, 896
gigas, 892, 893, 894
mackini, 891
Branchinella, 895, 897
sublettei, 888
Branchiobdella
astaci, 458
hexodonta, 457
kozarovi, 458
parasita, 458
Branchiocaris, 867
Branchiura, 10, 436, 438, 443, 453
sowerbyi, 439, 445, 452
Bratislavia, 451
Bresslaua, 89
vorax, 88
Brillia, 690
Bryocamptus, 921, 938
arcticus, 950
calvus, 950
cuspidatus, 950
douwei, 950
hiatus, 950
hiemalis, 950
hutchinsoni, 950
minisculis, 950
minnesotensis, 950
minutus, 950
morrisoni, 950
newyorkensis, 950
nivalis, 950
pilosus, 950
pygmaeus, 950
1040
Taxonomic Index
Bryocamptus (continued)
subarcticus, 950
tikchikensis, 950
umiatensis, 950
vejdovskyi, 950
washingtonensis, 950
zschokkei, 950
Bryochoerus, 542
Bryocyclops, 936
muscicola, 949
Bryodelphax, 542
parvulus, 530
Bryospilus, 847, 874
Buddenbrockia, 513
Buenoa, 678, 681
Bulimnea megasoma, 320, 321
Bulinus globosus, 300
Bullatifrons, 874
Bunops, 883
acutifrons, 885
Bursaria, 62, 83
truncatella, 84
Byrophyllum, 85
lieberkÅhni, 84
Bythotrephes, 852, 859, 862, 865, 887
cederstroemii, 886, 927
longimanus, 887
Cactus, 883
Caecidotea, 14, 785, 786, 796
recurvata, 788, 791
tridentata, 788, 791
Caenestheria, 900
setosa, 892
Caenestheriella setosa, 899
Caenis, 662, 698
Caenomorpha, 83
medusula, 82
Caenorhabditis briggsae, 263
Calcarobiotus, 529, 531, 532
Calineuria, 740, 760
californica, 756
Calliasmata, 961
Callibaetis, 662
Callinectes, 459
sapidus, 960
Calliobdella vividia, 481
Calohypsibius, 529, 531, 532, 543
Calopteryx, 701
Calposoma, 6, 135, 148, 151, 153
dactyloptera, 148
Calyptotricha pleuronemodies, 86
Cambarellus, 963, 964, 965, 967, 968, 972,
983, 988
puer, 985
Cambarincola, 460
fallax, 461
Cambarus, 821, 963, 964, 966, 967, 968,
983, 989
aculabrum, 982
alleni, 974
bartonii, 955, 981, 983
brachydactylus, 960
cahni, 960
chaugaensis, 974
diogenes, 957, 985
distans, 957
friaufi, 960
gentryi, 957
laevis, 960
latimanus, 974, 987
ornatus, 960
robustus, 956, 972, 974, 983
strigosus, 953
tenebrosus, 956, 960
zophonastes, 983
Camelobaetidius, 662
Campanella, 81
umbellaria, 80
Campascus, 76
triqueter, 76
Campbellonemertes, 174
Campeloma, 300, 303, 309, 323
decisum, 304, 305, 309, 324
rufrum, 305
Campostoma anomalum, 782
Camptocercus, 874, 875
liljeborgi, 876
macrurus, 876
oklahomensis, 876
rectirostris, 875, 876
Candocypria, 830, 835
Candocyprinotus, 830, 832
Candona, 14, 830, 832
acuta, 821
acutula, 820
anceps, 821
candida, 816, 818, 819, 820, 822
caudata, 809, 814, 819, 821
decora, 819, 822
ikpikpukensis, 821, 829
jeanneli, 821
marengoensis, 821
mulleri, 821
ohioensis, 820, 824
paralapponica, 821
patzcuaro, 822, 823
pedata, 821
protzi, 818, 821
rawsoni, 822, 827, 831
rectangulata, 817
reducta, 816
renoensis, 821
subtriangulata, 815, 817, 819, 820, 821,
822, 824
suburbana, 811
Candonocypris
deeveyi, 829
novaezelandiae, 813, 822
pugionis, 829
sarsi, 829
serrato-marginata, 829
Canthocamptus, 921, 938
assimilis, 950
oregonensis, 950
robertcokeri, 950
sinuus, 950
staphylinoides, 950
staphylinus, 950
vagus, 950
Capitomermis, 280
Capnia, 730
Carchesium, 81
polypinum, 80
Caridinophila, 458
Carphania, 529, 530, 531, 532, 542
fluviatilis, 538
Castrada, 157, 162, 170
hofmanni, 180
lutheri, 180
virginiana, 171, 180
Castrella, 173
graffi, 179
pinguis, 171, 179
Catenula, 157, 163, 165
confusa, 179
lemnae, 158, 165, 179
leptocephala, 165, 179
sekerai, 179
turgida, 179
virginia, 179
Caudatella, 662
Cavernocypris, 830, 832
subterranea, 822
wardi, 822
Celina, 683
Cenocorixa, 678
bifida, 575
Centrolimnesia, 555, 643
bondi, 617
Cephaliophora, 229
Cephalobus, 274
Cephalodella, 197, 202, 204, 245, 247
forficula, 232
Cephalothamnium, 52, 71
cyclopum, 71
Ceraclea, 672, 705
Ceratium
furcoides, 931
hirudinella, 46
Ceratodrilus, 10, 460
thysanosomus, 460
Ceratophylum demersum, 306, 307
Ceratopsyche, 672
Cercobrachys, 662
Cercomonas, 52, 71, 72
Cercopagis, 887
pengoi, 887
Ceriodaphnia, 852, 855, 856, 857, 857,
859, 861, 864, 867, 881
quadrangula, 880
Chaena, 85
teres, 84
Chaetogaster, 437, 439, 450
diaphanus, 824
limnaei, 384
Chaetonotus, 7, 185, 188, 189, 190
Chaetospira, 82
mülleri, 82
Chaoborus, 14, 34, 205, 225, 690, 715,
730, 731, 852, 857, 863, 920, 927, 929
Taxonomic Index
Chaoborus flavicans, 824
Chaos, 74
illinoisense, 73
Chappuisides, 559, 566, 643
eremitus, 626
Chara, 816, 823
fragilis, 823
Chauliodes, 677
Cheiroseius, 632
Chelohydracarus, 559, 644
navarrensis, 627
Chelomideopsis, 559, 644
besselingi, 627
Chilodonella, 55, 78
uncinata, 88
Chilomonas, 71
paramecium, 66
Chimarra, 705
Chironomus, 13, 573, 716, 730, 731, 734,
735
riparus, 482
tentans, 735
Chlamydomonas, 69, 396
reinhardtii, 516
Chlamydomyxa, 77
montana, 76
Chlamydophrys, 75
minor, 75
Chlamydotheca, 830, 833
arcuata, 812, 820
Chlorella, 6, 62, 143, 187, 219, 396, 870
fusca, 229
vulgaris, 395
Chlorohydra, 6, 143, 151
Chordarium, 165
europaeum, 179
Chordodes, 288, 292
morgani, 282
Chordodiolus, 291
Chromadora, 265
Chromadorita, 276
Chromogaster, 247, 276
africana, 264
troglodytes, 260, 264, 277
Chrysops, 715, 730
Chydoris, 143, 857, 859, 874, 878
brevlabris, 879
faviformis, 869
piger, 878
reticulatus, 869
sphaericus, 865, 869, 878
Ciliophrys, 78
infusionum, 77
Cincinnatia, 320
Cinetochilum, 87
margaritaceum, 86
Cipangopaludina, 323
chinensis, 303
japonica, 324
Cladotanytarsus mancus, 581
Clappia, 320
Clathrosperchon, 554, 563, 579, 607, 639
Clathrostoma, 89
viminale, 88
Clathrulina, 77
elegans, 77
Cletocamptus, 937
albuquerquensis, 950
brevicaudata, 950
deitersi, 950
Climacia, 710
Climacostomum, 85
virens, 84
Cloeodes, 662
Cochliopina, 320
Cochliopodium, 74
bilimbosum, 75
Codosiga, 71
botrys, 72
Coenagrion puella, 575
Cohnilembus, 87
pusillus, 86
Coleps, 85
hirtus, 67, 84
Colletheca, 202, 214, 224, 235, 236
cornuta, 200
gracilipes, 213, 214
trilobata, 200
Colpidium, 89
colpoda, 57, 88
Colpoda steinii, 88
Columbiathyas, 553, 640
crenicola, 610
Colurella, 241
Colymbetes, 683
Condylostoma, 85
tardum, 84
Conocephalus, 730
Conochilioides, 212, 237
natans, 216
Conochilus, 211, 212, 213, 215, 227, 228,
237, 862
hippocrepis, 212
unicornis, 211
Cookacarus, 555, 642
columbiensis, 613
Cookidrilus, 443
Copelatus, 683
Coptotomus, 683
Coquillettidia, 691, 715
perturbans, 573, 580
Corbicula, 9, 354, 355, 397, 398, 399
fluminea, 332, 335, 336, 339 – 356,
359 – 363, 366 – 368, 370 – 377, 379,
381 – 392, 394 – 396, 398, 399
Cordulegaster, 701
Cordulia, 701
Cordylophora, 135, 148, 150, 151, 153
Coregonus autumnalis migratorius, 794
Cornechiniscus, 542
Coronarctus, 538
Corophium spinicorne, 780
Corticacarus, 556, 618, 647
Corvomeyenia, 111, 115, 128
carolinensis, 115, 117, 118, 129
everetti, 108, 111, 115, 118, 119, 129
Corvospongilla, 129
becki, 118, 129
novaeterrae, 118, 119, 129
Corydalus, 730
Cothurnia, 81
imberbis, 80
Cottus carolinae, 791
Cowichania, 554, 639
interstitialis, 608
Crangonyx, 785, 786, 787, 792, 796
Craspedacusta, 6, 135, 136, 139, 140,
146 – 148, 150, 151, 152, 153
sinensis, 147
sowberii, 146, 147
Creaseria, 961
Crenitis, 684
Cretacimermis, 262
libani, 262, 281
Cricotopus, 39, 264, 690
bicinctus, 754, 755
nostocicola, 750, 751, 756
sylvestris, 754, 755
Cristatella, 509, 510
magnifica, 520
mucedo, 511, 512, 513, 522, 523
Cronodrilus, 460, 461
Crustipellis, 435, 448
tribranchiata, 447
Cryphiops, 961
Cryptocyclops, 937
bicolor, 949
Cryptomonas, 50, 931
Cryptonchus, 278
Cubozoa, 145
Culex, 730
pipiens quinquefasciatus, 269
Culicimermis, 280
Cumberlandia monodonta, 404, 408
Cupelopagis, 199, 204, 224, 235
vorax, 236
Cura, 167
foremanii, 161
Cyclestheria, 887
hislopi, 890
Cyclidium, 87
glaucoma, 86
Cyclocypris, 829, 830, 835
ampla, 810, 818, 819, 820, 821,
824, 834
globosa, 821
laevis, 821
ovum, 816, 825
sharpei, 818, 819, 820, 821
Cyclomomonia, 558, 643
andrewi, 625
Cyclonaias tuberculata, 404, 409
Cyclops, 921, 936
canadensis, 949
columbianus, 949
furcifer, 949
kolensis, 949
laurenticus, 949
scutifer, 921, 949
singularis, 922
strenuus, 949
vicinus, 923, 928, 949
1041
1042
Taxonomic Index
Cyclothyas, 554, 640
rivularis, 609
siskiyouensis, 609
Cylindrolaimus, 276
Cymatia bonsdorfi, 580
Cymbella, 974
Cymbiodyta, 684
Cymocythere, 830, 837
Cyphoderia, 76
ampulla, 76
Cyphon, 713
Cypretta, 830, 832
kawatai, 823
Cypria, 816, 822, 824, 830, 835
obesa, 821
ophtalmica, 818, 819, 820, 825
reptans, 813
turneri, 810, 814, 823
Cypricercus, 812, 823, 828, 829, 830, 833
deltoidea, 814, 821
fuscatus, 814
horridus, 814, 823
reticulatus, 814, 831
splendida, 814
tincta, 814
Cypriconcha, 828, 829
Cyprideis, 813, 816, 817, 830, 836
torosa, 816, 819
Cypridopsis, 823, 825, 829, 830, 832
hartwigi, 823
vidua, 809, 813 – 820, 823 – 826, 831
Cyprinotus, 816, 825, 828, 829, 830, 835
carolinesis, 814, 815, 816, 823
glaucus, 810, 814, 818
incongruens, 813, 814, 815, 823, 825, 826
salinus, 817
Cyprinus carpio, 385
Cypris, 823, 830, 835
balnearia, 820
bispinosa, 817
pubera, 824, 826, 834
reptans, 825
Cyprogenia, 409
stegaria, 404
Cyprois, 832
marginata, 831
Cyrenoida floridana, 397
Cyrtolophosis, 87
mucicola, 86
Cyrtonaias, 402, 404
tampicoensis, 416
Cyrtonia, 243
Cystobranchus, 490, 494
mammillatus, 494
meyeri, 468, 494
verrilli, 468, 494
virginicus, 494
Cytherissa, 826, 830, 836
lacustris, 815, 819, 821, 837
Cytheromorpha, 826, 830, 836
fuscata, 816, 817, 818, 819, 821
Cyzicus, 889, 891, 892, 900
californicus, 891, 899
setosa, 891
Dactylobiotus, 529, 531, 532, 533, 539,
543
dispar, 535
Dactylocythere, 830, 837
leptophylax, 837
Dadaya, 874, 878
macrops, 877
Dalyellia, 162, 164, 173
viridis, 158, 179
Daphnella, 859
Daphnia, 13, 15, 64, 150, 205, 226, 780,
847, 850 – 857, 859 – 867, 880, 881,
890, 924, 926 – 929
catawba, 867
cuculata, 867
denticulatus, 859
galeata, 852, 867
jollyi, 867
laevis, 864
longiremis, 867
longispina, 855
lumholtzi, 853, 858
magna, 853, 855, 857
pulex, 226, 854, 856, 859, 861, 864, 926
pulicaria, 848, 849, 852, 864
retrocurva, 849, 852, 867
similis, 39
Daphniopsis, 854, 880, 881
ephereralis, 855, 880
Darwinula, 810, 813, 814, 826, 829, 830
stevensoni, 811, 815, 831
Desydytes, 191
ornatus, 187
Dasyhormus, 165
Daubaylia, 261, 274, 275
Deltopylum, 67
Dendrocoelopsis, 157, 169
Dendrocometes, 81
paradoxus, 79
Dendrosoma, 79
radians, 79
Dero, 436, 438, 439, 446, 451, 452
Desmarella, 71
monoliformis, 72
Desmolaimus, 265
Desmona, 672
Desmopachria, 683
Desserobdella, 488, 492
michiganensis, 493
phalera, 493
picta, 493
Deuterophlebia, 716
Dexteria, 897
floridanus, 896
Diacyclops, 917, 921, 936
alabamensis, 949
albus, 949
bernardi, 949
bicuspidatus, 949
bisetosus, 949
chrisae, 949
crasscaudis, 949
dimorphus, 949
harryi, 949
haueri, 949
hypnicola, 949
jeanneli, 949
languidoides, 949
languidus, 949
nanus, 949
nearcticus, 949
palustris, 949
sororum, 949
thomasi, 921, 949
yeatmani, 949
Diacypris
compacta, 823
dietzi, 823
Diamphidaxona, 556, 566, 616, 643
Diaphanosoma, 851, 852, 857, 874
birgei, 872
Diaptomus, 935
glacialis, 948
connexus, 39
Dibusa angata, 752
Dicamptodon ensatus, 756
Dichaetura, 185, 188, 189, 190
Dicosmoecus, 672, 739, 749, 760
gilvipes, 739, 743, 750, 756
Dicranophorus, 202, 239
Dicrotendipes, 690
Dictya, 690
Didinium, 48, 49, 51, 62, 85
nasutum, 84
Difflugia, 67, 74
corona, 75
Dileptus, 62, 85
anser, 86
Dina, 487, 489, 496
anoculata, 497
dubia, 497
parva, 497
Dineutus, 684, 713
Dinobryon, 71
sertularia, 66
Dioctophyme renale, 459
Diphascon, 529, 531, 532, 533, 534, 539,
544
Diphetor, 662
Diploeca, 71
placita, 72
Diplois daviesiae, 241
Diploperla, 669
Diplophyrys, 77
archeri, 76
Discomorphella, 83
pectinata, 82
Disconaias, 402, 416
discus, 404
Discophyra, 81
elongata, 79
Disematostoma, 89
bütschlii, 88
Disparalona, 874, 878
acutirostris, 878
dadayi, 878
leei, 878, 879
rostrata, 878
Taxonomic Index
Dissotrocha, 235
Distocambarus, 963, 964, 966, 967, 983,
989
crockeri, 988
Dixella, 716
Dochmiotrema, 173
Dolania, 698
Dolerocypris, 830, 833
Dolomedes, 12, 550, 632, 649
scriptus, 649
striatus, 649
tenebrosus, 649
triton, 649, 650
vittatus, 649
Donacia, 713
Donnaldsoncythere, 830, 837
Dorylaimus, 257, 262, 265, 271, 274, 275
atratus, 263
Doryphoribius, 529, 531, 532, 539, 543
Dosilia, 128
palmeri, 119, 128
radiospiculata, 119, 120, 128
Dracunculus medinensis, 929
Dreissena, 9, 360, 397
bugensis, 332, 343, 350, 356, 360
polymorpha, 113, 332, 338 – 340, 343,
345 – 350, 352 – 356, 358 – 363,
366 – 368, 372 – 373, 375, 376, 378,
381 – 397, 929, 975
Drepanothrix, 883
dentata, 884
Drepanotrema, 318
kermatoides, 317
Drilomermis, 280
Dromogomphus, 666
Dromus dromas, 404, 410
Drunella, 662, 760
Dubiraphia, 684
Dugesia, 6, 163, 167
dorotocephala, 168
polychroa, 158, 168
tahitienseis, 161
tigrina, 157, 159, 168
Dunhevedia, 874, 878
americana, 877
Duosclera, 120
mackayi, 98, 106, 120, 121, 129
Dysnomia triquetra, 396
Dytiscus, 288, 683
Echinamoeba, 74
exudans, 73
Echiniscoides, 529, 538
sigismundi, 536
Echiniscus, 529, 531, 532, 538, 541, 542
spiniger, 528
Echinursellus longiunguis, 538
Eclipidrilus, 448, 449, 450
Ectemnia, 690
Ectocyclops, 936
phaleratus, 949
polyspinosus, 949
rubescens, 949
Ectopria, 713
Eiseniella, 446
Elaphoidella, 938
amabilis, 950
bidens, 950
californica, 950
carterae, 950
kodiakensis, 950
reedi, 950
shawangunkensis, 950
subgracilis, 950
tenuicaudis, 950
wilsonae, 950
Elgiva, 690
Elimia, 304, 305, 307, 310, 314, 323
alabamiensis, 314
caelatura, 314
cahawbensis, 309
carinocastata, 314
crenatella, 314
cylindracea, 314
fascians, 314
gerhardtii, 314
haysiana, 314
hydei, 314
livescens, 322
olivula, 314
showalteri, 314
Ellipsaria, 402
lineolata, 404, 410, 414
Elliptio, 402, 403, 408, 411, 412, 414, 415
complanata, 348, 349, 359, 360, 367,
368, 381, 387, 390, 408
crassidens, 405
dilatata, 412
icterina, 408
lanceolata, 408
nigella, 357
Elliptoideus, 402, 405
sloatanus, 413
Ellisodrilus, 458, 460
Elodea, 213, 214, 231, 986
canadensis, 213
Elodium, 150
Elosa, 244
Empidomermis, 280
Enallagma, 666, 701, 753, 754
clausum, 39
ebrium, 580
Encentrum linnhei, 232
Enchelyodon, 85
elegans, 84
Enchelys, 85
simplex, 84
Endochironomus, 690
Endotribelos, 690
Enochrus, 684
Entocythere, 776, 829, 830, 837
donnaldsonensis, 821
Entosiphon, 71
sulcatum, 66
Eocyzicus, 900
concavus, 899
Eogelyella carolinensis, 934
1043
Eohypsibius, 529, 531, 532, 539, 543
Eothinia, 202
Epactophanes, 938
richardi, 950
Ephemera, 698
Ephemerella, 662, 754
Ephemoroporus, 869, 874, 878
acanthodes, 877
Ephoron, 698
Ephydatia, 117, 130
fluviatilis, 101, 106, 107, 108, 110, 117,
120, 121, 130
millsii, 121, 130
müelleri, 99, 105, 106, 107, 108, 120,
121, 122, 130
Ephydra, 38, 730
Epioblasma, 402, 411, 414
aracaeformis, 357
biemarginita, 357
flexuosa, 357, 405
florentina, 357
haysiana, 357
lenior, 357
lewisii, 357
obliquata, 357
personata, 357
propinqua, 357
sampsonii, 357
stewardsonii, 357
torulosa, 357
turgidula, 357
Epiphanes, 202, 228, 242, 243
senta, 222
Epischura, 227, 912, 919, 922, 925, 926,
934
baikalensis, 911
fluviatilis, 948
lacustris, 921, 923, 948
massachusettsensis, 948
nevadensis, 948
nordenskioldi, 948
Epistylis, 81
plicatilis, 80
Epitheca, 744
cynosura, 743
princeps, 743
simplicollis, 746
Erebaxonopsis, 558, 647
nearctica, 622
Eremobiotus, 529
Ergasilus
arthrosis, 950
auritus, 950
caeruleus, 950
celestis, 950
centrarchidarum, 950
cerastes, 950
chautauquaensis, 950
clupeidarum, 950
cotti, 950
cyprinaceus, 950
elongatus, 950
japonicus, 950
lanceolatus, 950
1044
Taxonomic Index
Ergasilus (continued)
luciopercarum, 950
megaceros, 950
nerkae, 950
rhinos, 950
sieboldi, 950
tenax, 950
versicolor, 950
wareaglei, 950
Eristalis, 715
Erpetocypris barbatus, 829
Erpobdella, 467, 483, 489, 496
octoculata, 474, 480, 482, 483
punctata, 472 – 476, 479, 480, 482 – 485,
496
Erythemus simplicicolis, 580
Escherichia coli, 382
Esox lucius, 782
Espejoia, 67
Estellacarus, 558, 637, 646
unguitarus, 599, 622, 623
Estelloxus, 555, 636, 642
montanus, 615
Ethmolaimus, 263, 276
Eubosmina, 857, 857, 881, 882
coregoni, 882
Eubranchipus, 15, 889, 891, 895, 897
bundyi, 893
holmni, 893
hundyi, 896
moorei, 891
serratus, 892, 893
vernalis, 893
Eubrianax, 687
Eucephalobus, 274
Euchlanis, 198, 204, 223, 241, 242
dilatata, 205, 209, 217, 232
Euchordodes, 292
nigromaculatus, 283, 287, 288, 289
Euchromadora striata, 263
Eucyclops, 921, 936
agilis, 949
bondi, 949
conrowae, 949
elegans, 949
macruroides denticulatus, 949
neomacruroides, 949
prionophorus, 949
Eucypris, 822, 823, 828, 830, 833
crassa, 834
serrata, 828
serrato-marginata, 829
virens, 823
Eudiaptomus, 935
gracilis, 928, 931, 948
Eudorylaimus andrassyi, 263
Euglena, 63
Euglypha, 76
tuberculata, 76
Eukiefferiella, 690
Eulimnadia, 897
agasszii, 898
antelei, 890, 892
diversa, 890
inflecta, 891
Eunapius
fragilis, 106, 120, 121, 122, 130
igloviformis, 120
mackayi, 120
Eupera, 400
Euplotes, 56, 62, 81
patella, 58, 80
Euryalona, 874, 876
orientalis, 877
Eurycercus, 853, 859, 862, 869, 874
lamellatus, 875
Eurylophella, 662, 699
Eurytemora, 934
affinis, 948
americana, 948
arctica, 948
bilobata, 948
canadensis, 948
composita, 948
gracilicauda, 948
Euteratocephalus, 276
Euthyas, 554, 633, 640
mitchelli, 609
truncata, 609
Evadne, 857, 859, 887
nordmanni, 886
Eylais, 553, 568, 570 – 577, 579, 580, 581,
607, 631, 634, 639
discreta, 569, 570
eurylhalina, 569
major, 571
Fabaeformiscandona, 809, 814, 818, 819,
821, 830, 833
pennaki, 822
wegelini, 817, 822
Fallceon, 662
Fallicambarus, 963, 964, 966, 967, 983,
989
devastator, 974, 976, 985
fodiens, 977, 988
gordoni, 977
Farula, 676
Faxonella, 963, 964, 965, 989
clypeata, 981, 985
creaseri, 987
Feltria, 557, 577, 593, 631, 637, 643
exilis, 617
Ferrissia, 316
fragilis, 310
rivularis, 317
Filamoeba, 74
nolandi, 73
Filinia, 205, 208, 209, 227, 237
longiseta, 221, 227
Fisherola, 316
nutalli, 317
Flabelliforma ostracodae, 825
Floscularia, 211, 212, 214, 224, 237
conifera, 211, 212, 213, 214, 219, 238
Fluminicola, 303, 320
Fontelicella, 320
Fontigens, 320
Forelia, 557, 566, 577, 579, 638, 645
floridensis, 620
onondaga, 596
ovalis, 596
Fossaria, 302, 320
humilis, 321
modicella, 304
Fragilaria, 974
Fredericella, 505, 506, 507, 510, 512, 513,
514, 517
browni, 512, 518, 519
indica, 511, 511, 512, 518, 519, 522
sultana, 511, 518
Frontipoda, 555, 636, 641
americana, 615
Frontipodopsis, 557, 566, 579, 617, 643
Frontonia, 67, 89
leucas, 88
Fujiscon, 529, 531
Fusconaia, 402, 411, 414 flava, 361, 405
ebena, 385
masoni, 408
Gammarus, 14, 163, 785, 787, 796
lacustris, 782, 786, 790, 791, 792, 919
minus, 781, 782, 786, 787, 788, 791, 793
pulex, 785, 790
Gasterosteus aculeatus, 824
Gastropus, 247, 248
Gastrostyla, 81
steini, 82
Gatromermis, 278
Geayia, 559, 563, 579, 628, 635, 644
Geocentrophora, 157, 164, 166
cavernicola, 179
sphyrocephala, 179
Geocythere, 830, 837
Gerris, 570, 678, 709, 730
comatus, 580
Giardia, 69
Gieysztoria, 157, 173
Glaucoma, 89
scintillans, 88
Glebula, 402
rotundata, 405, 415
Gleotrichia, 238
Glossiphonia
complanata, 472, 476, 477, 479, 480,
482, 483, 488, 492, 494
heteroclita, 488
rudis, 488
Glossosoma, 706, 730, 737, 740, 760
Glypotendipes paripes, 482
Goera, 672, 706
Goerita, 672
Gomphaeschna, 666
Gomphonema, 974
Gomphus, 701, 730
Gonidea, 402
angulata, 403, 405
Gonostomum, 83
affine, 82
Taxonomic Index
Gordius, 282, 283, 285, 288, 291, 292
aquaticus, 282, 283
diblastus, 287
dimorphus, 283, 287
robustus, 288
tenuifibrosus, 280
Graphoderus, 683
Graptoleberis, 857, 874, 876
testudinaria, 862, 875
Grimaldina, 883
brazzai, 885
Guernella, 883
raphaelis, 885
Gulcamptus, 938
alakaensis, 950
huronensis, 950
laurentiacus, 950
Guttipelopia, 690
Gymnodinium, 66, 71
Gymnopais, 690
Gyratrix, 157, 173
hermaphroditus, 161, 169, 180
Gyraulus, 318
circumstriatus, 304
deflectus, 304, 310, 317
Gyrinus, 684, 730
Gyrodinium, 66, 71
Gyrotoma, 314
Haber, 453
speciosus, 452
Habrophlebia, 699
Habrotrocha, 235
rosa, 207, 231, 232
Haementaria, 488
ghilianii, 469, 474, 481
Haemonais waldvogeli, 451
Haemopsis, 489
sanguisuga, 480
Halicyclops, 914, 934
Halipegus occidualis, 825
Haliplus, 684, 711, 713
Halobates, 742
Halobiotus, 529, 537
crispae, 535
Haltdytes, 191
Halteria, 62, 83
grandinella, 82
Haplohexapodibius, 529
Haplomacrobiotus, 529, 531, 532,
533, 543
Haploperla, 702
Haplotaxis, 435, 437, 439, 441, 446, 447
Haptoglossa, 229
Harpagocythere, 830, 838
Harringia, 239, 240
Hartmanella, 74
vermiformis, 73
Hartocythere, 830, 837
Hastatella, 81
radians, 80
Hauffenia, 320
Hebesuncus, 529, 531, 544
Hebetancylus, 316
excentricus, 317
Heleidomermis, 280
Heleidomermis libani, 262
Helichus, 686, 712
Helicopsyche, 705, 760
borealis, 743, 750
Heliophyra, 81
reideri, 79
Helisoma, 307, 311, 318, 975
anceps, 304, 305, 308, 313, 319, 825
companulatum, 304
trivolvis, 302, 306
Helobdella, 469, 492
california, 493
elongata, 490, 493
fusca, 493
papillata, 493
robusta, 493
stagnalis, 472, 473, 476, 477, 479, 480,
482, 483, 492, 493
transversa, 493
triserialis, 479, 493
Helophorus, 688, 712
Helopicus, 669
Hemicycliophora, 270, 273
Hemigrapsis estellinensis, 960
Hemistena, 402
lata, 405, 411
Heptagenia, 662
Herpetocypris, 815, 816, 830, 834
brevicaudata, 808
reptans, 816, 824
Hesperocorixa, 678
Hesperodiaptomus, 912, 927, 929, 935
arcticus, 948
augustaensis, 948
breweri, 948
caducus, 948
californensis, 948
eiseni, 948
franciscanus, 948
hirsutus, 948
kenai, 948
kiseri, 948
nevadensis, 923, 948
novemdecimus, 948
schefferi, 948
septentrionalis, 926
shoshone, 926, 931, 948
victoriaensis, 948
wardi, 948
Hesperoperla, 669
Hesperophylax, 672, 705
Hetaerina, 701
Heterocope, 912, 925, 934
septentrionalis, 922, 926, 948
Heterocypris, 823, 828, 829
incongruens, 814, 815
Heterolepidoderma, 187, 188, 190
Heteromeyenia, 117, 129
baileyi, 121, 122, 123, 124, 129
latitenta, 122, 123, 124, 129
tentasperma, 122, 123, 124, 129
1045
tubisperma, 122, 123, 124, 129
Heteromita, 52
Heterophrys, 78
myriopoda, 77
Heteroplectron, 706
californicum, 748, 749, 756
Heterosternuta, 683, 684
Hetrophrys, 50
Hexagenia, 698, 730, 732, 735
Hexamita, 52, 57, 71
inflata, 72
Hexapodibius, 529, 531, 532, 543
Hexarthra, 203, 205, 227, 236
fennica, 39, 216, 221
Hexatoma, 715
Heydenius dominicanus, 262, 281
Hirschmanniella, 270
Hirudo medicinalis, 469, 470, 474, 481,
482, 486, 496
Histobalantium, 87
natans, 86
Hobbseus, 963, 964, 966, 967, 983, 989
cristatus, 988
Holopedium, 857, 859, 861, 863, 864
amazonicum, 873
gibberum, 872, 873
Holophyra, 85
simplex, 86
Homalozoon vermiculare, 84
Homocaligus muscorum, 648
Homochaeta naidina, 450
Homocyclops, 936
ater, 949
Homoeoneuria, 662, 699
Hoplodictya, 690
Horatia, 320
Horreolanus, 559, 645
orphanus, 628
Hoyia, 320
Huitfeldtia, 557, 579, 638, 648
rectipes, 573, 595, 619
Huntemannia, 937
lacustris, 950
Hyalella, 793, 794, 796
azteca, 483, 781, 786, 787, 790 – 793
montezuma, 477, 483, 786, 790
Hyalinella, 514
orbisperma, 511, 521
punctata, 511, 513, 514, 519, 521
Hyalopyrgus, 320
Hyalosphenia, 74
cuneata, 75
Hydaticus, 683
Hydatophylax, 740
Hydra, 6, 136, 137, 138, 140 – 146, 150,
151, 152, 754, 755, 850
attenuata, 145
braueri, 152, 153
oligactis, 152, 153
viridissima, 151, 153
vulgaris, 152, 153
Hydrachna, 553, 568, 569, 572 – 575, 579,
580 – 582, 606, 631, 634, 639
conjecta, 570, 580
1046
Taxonomic Index
Hydrachna (continued)
magniscutata, 587
virella, 580
Hydramoeba hydroxena, 140
Hydrocanthus, 684, 712, 713
Hydrochara, 684
Hydrochoreutes, 557, 638, 647
intermedius, 595, 619
minor, 595
Hydrochus, 712
Hydrodroma, 554, 579, 581, 633, 639
despiciens, 568, 571, 581
Hydrolimax, 166
grisea, 157, 179
Hydromermis, 280
Hydrometra, 709
Hydroperla, 669
Hydroporus, 683, 684, 711, 730
Hydropsyche, 672, 730, 739, 740, 757, 760
Hydroptila, 705
Hydroscapha, 712
natans, 682
Hydrovatus, 684
Hydrovolzia, 553, 569, 570, 571, 633, 638
gerhardi, 563
marshallae, 606
Hydrozetes, 632, 648
Hydrozoa, 145
Hydryphantes, 553, 564, 572, 579, 581,
608, 633, 639
ruber, 565, 608
tenuabilis, 568, 572, 574
Hygrobates, 556, 569, 572, 574, 579, 582,
593, 618, 637, 647
fluviatilis, 582
neocalliger, 593
nigromaculatus, 581
Hygrotus, 684
Hyla regilla, 285, 288
Hylodiscus, 74
rubidicundus, 73
Hymanella, 168
retenuova, 168
Hypechiniscus, 529, 538, 542
Hypotrichidium, 83
conicum, 82
Hypsibius, 529, 531, 532, 534, 537, 539,
541, 543
convergens, 539
dujardini, 539
Ichthydium, 188, 189, 190
Ichthyophthirius, 67
Ictalurus
furcatus, 385
punctatus, 385
Ictiobus
bubalus, 385
niger, 385
Illinobdella, 468, 489, 490, 494
alba, 495
elongata, 495
richardsoni, 495
Ilybius, 684
Ilyocryptus, 861
sordidus, 884
Ilyocypris, 822, 830, 835
bradyi, 821
gibba, 819, 821
Ilyodrilus, 454, 455
Ilyodromus, 830, 833
Io, 323
fluvialis, 315, 322
Ironus, 265, 278
Ischnura, 666
verticalis, 580
Isochaetides, 453, 454
Isocypris, 830, 833
longicomosa, 829
Isogenoides, 669
Isohypsibius, 529, 531, 532, 533, 534, 537,
538, 539, 543
granulifer, 537
Isomermis, 280
Isoperla, 669, 702, 730
Itaquascon, 529, 531, 533, 534, 544
Itocyclops, 937
yezoensis, 949
Itura, 202
bostoniensis, 216
Janicea, 961
Javalbia, 558, 622, 646
Juga, 307, 323, 739, 740, 756, 760
Kawamuracarus, 555, 616, 642
Kellicottia, 201, 243, 244
Kenkia, 168
Keratella, 8, 198, 201, 205, 210, 226, 227,
243, 245
americana, 205, 217
cochlearis, 222, 225, 226, 227, 232, 244,
245
crassa, 210, 222
earlinae, 222
quadrata, 208, 221, 245
slacki, 227
tauracephala, 206
testudo, 205, 220, 226, 227, 232, 245
Kerona, 140
Khawkinea, 5, 71
halli, 66
Kijanebalola, 190, 191
Kincaidiana
freidris, 449, 450
hexatheca, 448, 449
Koenikea, 556, 579, 618, 637, 645
marshallae, 594
wolcotti, 630
Kongsbergia, 557, 624, 629, 646
Krendowskia, 559, 635, 644
similis, 604
Krumbachia, 169
hiemalis, 171, 180
minuta, 180
virginiana, 180
Kurzia, 874, 876
latissima, 876
longirostris, 875, 876
Labiobaetis, 662
Laccobius, 684
Laccophilus, 684
Laccornis, 684
Lacinea mobilis, 785
Lacinularia, 211, 212, 214, 227, 237
flosculosa, 199
Lacrymaria, 85
olor, 84
Ladona, 666
Laevapex, 316
fuscus, 304, 310, 317
Lambornella, 67, 68
Lamellidens
corrianus, 341, 347, 351, 354
marginalis, 391
Lampsilis, 342, 364, 365, 402, 408, 411,
412, 415
binominata, 357
claibornesis, 353
ovata, 365, 405
perovalis, 364, 365
radiata, 345, 346
siliquoidea, 345, 358, 359
teres, 358
Lanceimermis, 278
Lanthus, 666
Lanx, 316
patelloides, 317
Lara avara, 748
Lasmigona, 402, 408, 409, 410, 413
complanata, 410
costata, 405, 409
Lathonura, 883
rectirostris, 884
Latona, 852, 859, 873
setifera, 872
Latonopsis, 874
occidentalis, 872
Laversia, 559, 569, 578, 579, 636, 645
berulophila, 603, 604, 627
Lebertia, 555, 572, 590, 615, 636,
642
Lecane, 210, 243
inermis, 232
lunaris, 242
quadridentata, 205, 217
Lecythium, 75
hyalinum, 75
Lemabadion, 87
magnum, 88
Lemanea australis, 752
Lemiox, 402, 405
rimosus, 409
Lemna, 682
Lepadella, 210, 241
Lepidocaris, 867
Lepidochaetus, 189, 190
Taxonomic Index
Lepidodermella, 188, 189, 190
squamata, 182, 186, 187
triloba, 187
Lepidostoma, 706, 763
Lepidurus, 890, 900
arcticus, 889
couessii, 899
lemmoni, 891
Lepomis
cyanellus, 782
gibbosus, 311
gulosus, 385
macrochirus, 385
microlophus, 311, 385
Leptistheria, 900
compleximanus, 898
Leptodea, 402, 408, 410, 414
fragillis, 405, 410, 414
Leptodiaptomus, 919, 935
ashlandi, 948
assiniboiaensis, 948
coloradensis, 948
connexus, 948
insularis, 948
judayi, 948
minutus, 921, 923, 931, 948
moorei, 948
novamexicanus, 948
nudus, 948
pribilofensis, 948
sicilis, 948
siciloides, 948
signicauda, 948
spinicornis, 948
trybomi, 948
tyrrelli, 922, 923, 948
Leptodora, 35, 147, 852, 859, 860, 861,
862, 885
kindtii, 885, 886
Leptohyphes, 662, 698
Leptophlebia, 699
cupida, 739, 743
Leptoxis, 314, 323
ampla, 314
picta, 314
praerosa, 314, 322
taeniata, 314
Lepyrium, 320
Lesquereusia, 74
spiralis, 75
Lestes, 667, 701, 730
Lethaxona, 557, 621, 646
Lethocerus, 680
Leuctra, 703
Lexathonella, 557, 647
parva, 621
Lexingtonia, 402
dolabelloides, 405, 411
subplana, 408
Leydigia, 852, 874, 876
acanthocercoides, 877
leydigi, 862
Libellula, 730
lydia, 745, 746
Lieberkuehnia, 77
wagnerella, 76
Ligumia, 402, 412, 415
recta, 381, 406
subrostrata, 348, 350, 351, 352
Limmenius, 529, 531, 532, 534, 542
Limnadia, 900
lenticuraris, 898
Limnesia, 556, 568, 572, 579, 605, 636,
643
maculata, 575
marshalliana, 592
undulata, 577
Limnias, 211, 214
melicerta, 238
Limnocalanus, 63
Limnocalanus, 912, 917, 922, 925, 934
johanseni, 948
macrurus, 917
Limnochares, 553, 568, 570, 572, 573, 576,
579, 607, 634, 639
americana, 569, 570, 586
aquatica, 570, 572, 573, 574, 575,
580
Limnocodium, 135, 147
victoria, 146
Limnocythere, 826, 830, 838
ceriotuberosa, 818, 819, 822
friabilis, 826
herricki, 819
itasca, 809, 810, 819, 837
liporeticulata, 821
sanctipatricii, 811
sappaensis, 818
staplini, 816, 818, 820, 822, 827
verrucosa, 818, 819
Limnodrilus, 439, 440, 445, 454, 455
cervix, 439
claparedeianus, 440
hoffmeisteri, 437, 440
profundicola, 438
Limnoithona, 936
sinensis, 949
Limnomermis, 280
Limnophora, 715, 678
Limnoruanis, 169
romanae, 159, 171, 180
Limnosida, 873
Limnozetes, 649
Limonia, 715, 730
Limpnephilus, 732
Linderiella, 897
occidentalis, 896
Lindia, 237, 240
Liodessus, 684
Lioplax, 303, 323
subcarinata, 324
Lioporeus, 684
Lipsothrix, 748
Lirceus, 785, 786, 796
fontinalis, 782, 791
usdagalun, 792
Lithasia, 323
geniculata, 322
1047
Lithocolla, 78
globosa, 77
Litocythere, 830, 838
Litonotus, 87
fasciola, 86
Ljania, 558, 638, 646
bipapillata, 598
michiganensis, 621
Lobohalacarus, 648
Lophocharis, 241
Lophopodella, 508, 510, 512
carteri, 510, 511, 512, 513, 516, 520,
523
Lophopus, 510
crystallinus, 511, 512, 520, 523
Lordocythere, 830, 836
Loxocephalus, 87
plagius, 86
Loxodes, 55, 56, 57, 85
magnus, 86
Loxophyllum, 51, 87
helus, 86
Lumbriculus, 438, 439, 445, 448, 449
variegatus, 438, 440, 445, 448
Lutrochus, 713
Lymnaea, 304, 307
elodes, 305, 307, 308, 311
emarginata, 305
peregra, 305, 306
stagnalis, 39, 298, 305, 314, 320, 321
Lynceus, 900
brachyurus, 898
Lyrogyrus, 310, 320
Macdunnoa, 662
Macrobdella, 495
decora, 469, 495
diplotertia, 495
ditetra, 495
sestertia, 495
Macrobiotus, 529, 531, 532, 534, 537, 539,
541, 543
areolatus, 538
hufelandi, 530, 537, 538, 539
persililis, 540
tonollii, 528, 538
Macrobrachium, 774, 783, 951, 961, 962,
963, 964, 988
acanthurus, 961, 962
carcinus, 961, 962
ohione, 783, 961, 962, 969, 970
olfersii, 961
rosenbergii, 959, 962
Macrochaetus, 243, 244
Macrocotyla, 168
Macrocyclops, 921, 929, 936
albidus, 949
fuscus, 912, 913, 949
Macrohectopus, 794
branickii, 790, 794
Macromia, 701
Macrostomum, 6, 157, 161, 166
collistylum, 179
1048
Taxonomic Index
Macrostomum (continued)
curvistylum, 179
gilberti, 167, 179
glochostylum, 179
lewisi, 179
lineare, 179
norfolkense, 179
ontarioense, 179
phillipsi, 179
reynoldsi, 179
riedeli, 179
ruebushi, 179
sensitivum, 179
sillimani, 179
stirewalti, 179
tennesseense, 179
tuba, 167, 179
virginianum, 179
Macrothrix, 859, 883
rosea, 885
Macrotrachella, 235
quadricornifera, 223
Macrovelia, 709
Macroversum, 529, 531, 532, 539
Magmatodrilus, 460
obscurus, 461
Malirekus, 669
Mamersellides, 555, 613, 642
Mansonia, 691
Maraenobiotus, 938
brucei, 950
canadensis, 950
insignipes, 950
Margaritifera, 366, 402, 412
falcata, 403
hembeli, 361
margaritifera, 350, 351, 358, 364, 366,
367, 368, 403, 404
Marinellina, 181, 182, 185, 188
flagellata, 181, 182
Marisa, 303, 318
cornuarietis, 324
Marstonia, 320
Martarega, 681
Marvinmeyeria, 469
lucida, 479, 491
Maryana, 89
socialis, 88
Mastigodiaptomus, 935
albuquerquensis, 948
texensis, 948
Matus, 684
Mayorella, 74
bigemma, 73
Medionidus, 402, 409, 413
conradicus, 406
mcglameriae, 357
Megacyclops, 937
donnaldsoni, 949
gigas, 912, 949
latipes, 949
magnus, 949
viridis, 921, 949
Megadrili, 446
Megafenestra, 881
nasuta, 880
Megalocypris, 828, 829, 830, 834
alba, 821, 823
deeveyi, 829
novaezelandiae, 813, 822
pugionis, 829
sarsi, 829
Megalonaias, 363, 402
nervosa, 406, 409, 413
Melampus, 313
Melanoides, 320
tuberculata, 322
Melanopsis, 314
Melosira, 516
Menetus dilatatus, 317
Meramecia, 556, 643
occidentalis, 616
Mesobates, 556, 618, 647
Mesochra, 938
alaskana, 950
Mesocrista, 529, 531, 532, 544
Mesocyclops, 227, 921, 929, 936
americanus, 913, 949
edax, 919, 920, 921, 922, 927, 928, 949
leuckarti, 917, 921
longisetus, 949
ogunnus, 917
ruttneri, 949
venezolanus, 949
Mesodinium, 85
pulex, 84
Mesodorylaimus, 274
Mesomermis, 280
Mesorhabditis, 274
Mesostoma, 157, 158, 159, 160, 161, 162,
163, 173
arctica, 161, 180
californicum, 161, 180
columbianum, 180
craci, 169, 172, 180
curvipenis, 180
ehrenbergi, 158, 159, 160, 161,
162, 180
lingua, 158, 159, 160, 162
macroprostatum, 180
platygastricum, 180
vernale, 161, 171, 180
virginianum, 180
Mesovelia, 709
Metacineta, 78
mystacina, 79
Metacyclops, 937
cushae, 916, 949
Metacypris, 830, 838
maracoensis, 821
Metacystis, 85
recurva, 84
Metopus, 83
es, 82
Metrobates, 678
Micractinium, 225
Micrasema, 672
Microbiotus, 529, 532
Microcometes, 77
paludosa, 76
Microcyclops, 937
pumilis, 949
rubellus, 949
varicans, 921, 949
Microdalyellia, 157, 173
abursalis, 179
circobursalis, 179
fairchildi, 179
gilesi, 179
rheesi, 179
rochesteriana, 179
rossi, 167, 179
ruebushi, 179
sillimani, 179
tennesseensis, 167, 179
virginiana, 179
Microgromia, 77
haeckeliana, 76
Microhydra ryderi, 146
Microhypsibius, 529, 531, 543
Microkalyptorhynchus, 173
Microlaimus, 274
Micropterus dolomieui, 979
Microrhynchus virginianus, 180
Microsporidium, 228
Microstomum, 157, 166, 265
lineare, 167, 179
Microthorax, 89
pusillus, 80
Midea, 552, 558, 577, 635, 644
expansa, 568, 602, 625
Mideopsis, 558, 566, 578, 579,
627, 645
borealis, 603
Mikracodides, 243
Milnesium, 529, 531, 532, 533, 534, 536,
539, 541, 542
Minibiotus, 529, 531, 532, 533, 543
Minytrema melanopus, 385
Mixibius, 529, 531, 532, 539
Mixodiaptomus, 935
theeli, 948
Moina, 850, 851, 852, 854, 857, 859, 861,
867, 881, 889, 890
wierzejskii, 886
Moinodaphnia, 881
macleayii, 886
Molanna, 672
Mollibdella, 489
grandis, 469, 496
Momonia, 558, 579, 635, 643
campylotiba, 601
projecta, 625
Monas, 71
Monhystera, 263, 265, 274
grelachii, 263
ocellata, 263
Monhystrella, 274
Monoattractides, 555, 614, 641
Monodella texana, 784
Mononchulus, 276
Mononchus, 262, 278
Taxonomic Index
Monoporeia, 789, 790, 796
affinis, 786, 789, 791
Monopylephorus, 437
Monosiga, 48, 71
ovata, 72
Monospilus, 847, 874
dispar, 875
Monostyla, 243
Mooreobdella, 467, 487, 489, 496
bucera, 497
fervida, 479, 497
melanostoma, 497
microstoma, 482, 497
tetragon, 497
Mopeschiniscus, 542
Moraria, 938
affinis, 950
arctica, 950
cristata, 950
duthiei, 950
laurentica, 950
mrazeki, 950
virginiana, 950
Morimotacarus, 559, 645
nearcticus, 626
Morone saxatilis, 385
Motobdella, 469, 470, 472, 483, 496
montezuma, 465, 477, 481, 483, 484,
489, 496
sedonensis, 484, 489, 497
Moxostoma carinatum, 385
Mucronothrus, 649
Multifasciculatum, 81
elegans, 79
Murrayon, 529, 531, 532, 539
Musculium, 343, 360, 362, 368, 383, 390,
400
lacustre, 369
partumeium, 343, 346, 347, 348, 350,
360, 369, 381, 388, 396, 400
securis, 351, 369, 387, 400
transversum, 377, 381, 382, 383, 384,
387, 389, 400
Mya arenaria, 353
Myelostoma, 83
flagellatum, 82
Myostenostomum, 165
tauricum, 166, 179
Myriophyllum, 150, 231
Mysis, 14, 225, 786, 789, 794, 796
littoralis, 786
relicta, 35, 786, 788, 789, 790
Mystacides, 672
Mytilina, 241
Mytilocypris henricae, 808, 814
Mytilopsis, 392, 397
leucophaeata, 397
Mytilus edulis, 347
Myzobdella, 489
lugubris, 492, 494
Naegleria, 73
fowleri, 54
gruberi, 53, 54
Nais, 440, 446, 452
elinguis, 440
Najadicola, 384, 557, 579, 638, 647
ingens, 581, 597, 620
Nannocandona, 830
faba, 822
Nannochloris oculata, 226
Nannopus, 937
palustris, 950
Nassula, 63, 67, 78
ornata, 80
Natricola, 320
Nautarachna, 557, 638, 645, 645, 648
muskoka, 597
queticoensis, 620
Navicula, 67, 974
Nebalia, 233
Nebela, 74
collaris, 75
Nebrioporus, 684
Necopinatum, 529, 531, 532, 542
Nectomena, 290, 291
munidae, 283
Nectopsyche, 672
Necturus maculosus, 363, 385
Nehalennia speciosa, 580
Nelumbo lutea, 511
Nemoura, 702, 703
Neoacarus, 559, 579, 636, 644
minimus, 626
occidentalis, 603
Neoatractides, 555, 614, 641
Neoaturus, 557, 624, 646
Neobosmina, 881, 882
tubicen, 882
Neobrachypoda, 558, 638, 646
ekmani, 598, 623
Neochordodes, 292
occidentalis, 284, 285, 286, 288,
289
Neochoroterpes, 662
Neodasys, 187
Neoechinorhynchus
prolixus, 825
rutili, 825
Neoephemera, 698
Neogossea, 190, 191
Neohermes, 760
Neolimnochares, 553, 570, 586, 607, 634,
639
Neomamersa, 556, 566, 616, 643
californica, 616
chihuahua, 616
Neomideopsis, 558, 644
suislawensis, 625
Neomysis mercedis, 786, 788
Neopalaemonon, 961
Neophylax, 676, 706, 730
aniqua, 750
Neoplanorbis, 318
Neoplea, 709, 732
Neoporus, 684
Neothremma, 676
Neothrix, 883
armata, 885
Neotiphys, 557, 638, 648
pionoidellus, 595, 619
Neotyrrellia, 556, 643
anitahoffmannae, 617
Nepa, 709
Nephelopsis, 483, 496
obscura, 312, 470 – 485, 487, 497
Neritina, 318
reclivata, 303, 319
Neumania, 556, 581, 618, 637, 647
papillator, 576, 577, 578
punctata, 594
Neurocordulia, 701
Niqronia, 677, 707
Nitokra, 937
hibernica, 950
lacustris, 950
Nordodiaptomus, 935
alaskaensis, 948
Nosema stenocypris, 825
Nostoc, 264
parmelioides, 750, 751, 756
Notholca, 204, 210, 244, 245
acuminata, 208
squamala, 245
Notoalona, 874, 876
freyi, 877
Notodromas, 830, 832
monacha, 813, 816
Notogilla, 320
Notommata, 201, 202, 215, 245, 247
copeus, 224, 232
Notonecta, 678, 681, 709, 857
Notopanisus, 554, 608, 640
Nüchterleinella corneliae, 825
Nudomideopsis, 558, 577, 635, 644
magnacetabula, 601, 602, 625
Nuphar, 511
Nyctiophylax, 672
Nygolaimus, 274
Nymphaea, 511
Nymphula, 710
Obliquaria, 402
reflexa, 378, 406, 410, 413
Obovaria, 402, 406, 411, 414
Ochromonas, 50, 71
variabilis, 66
Octomyomermis, 280
Octotrocha speciosa, 199
Odontolaimus, 274, 276
Odontomyia, 715
Oecetis, 672, 705
Oedipodrilus, 460
Okriocythere, 830, 836
Oligobdella, 469, 491
biannulata, 490, 492
Oligostomis, 705
Olisthanella, 157, 173
truncala, 180
Omartacarus, 556, 579, 616, 642
1049
1050
Taxonomic Index
Onchobunops, 883
Oncholaimus, 276
Oncorhynchus clarki, 756
Onychocamptus, 937
mohammed, 950
Onychodiaptomus, 935
birgei, 948
Opercularia, 81
nutans, 80
Ophidonais, 450, 451
serpentina, 450
Ophiogomphus, 666
Ophrydium, 81
eichhorni, 80
Ophryoglena, 59, 67
Ophryoxus, 883
gracilis, 884
Opisthocystis, 173
goettei, 169, 180
Opistomum, 169
pallidum, 158, 167, 180
Optioservus, 684
Orconectes, 963, 964, 966, 967, 968, 972,
983, 988, 989, 989
australis, 960, 966
compressus, 960
immunis, 955, 956, 972
inermis, 32, 959, 972, 987
luteus, 973, 974
nais, 955
neglectus chaenodactylus, 975
pellucidus, 956, 959
propinquus, 778, 958, 959, 972, 974,
975, 978, 979, 981, 982, 987
punctimanus, 973, 974
rusticus, 312, 955, 972, 974, 975, 976,
977, 978, 982, 985, 988
shoupi, 983
testii, 32
virilis, 778, 781, 959, 972, 976, 977, 978,
980, 981, 982, 983
Oreela, 529, 542
Oregonacarus, 555, 642
rivulicolus, 613
Oreodytes, 684
Ornamentula, 185, 191
Ornithocythere, 830, 836
Orohermes, 740
Orthocladius, 749
Orthocyclops, 936
modestus, 949
Ortmannicus, 967
Orygocerus, 320
Oscillatoria, 67
Osphranticum, 921, 935
labronectum, 948
Otomesostoma, 157, 166
auditivum, 160, 179
Oxus, 555, 572, 579, 615, 636, 641
elongatus, 591
Oxyethira, 705
Oxytrcha, 81
fallax, 82
Oxyurella, 874, 876
brevicaudis, 848, 877
Pacifastacus, 954, 963, 964, 966, 967, 983,
988
chenoderma, 966
fortis, 967, 983
gambelii, 963
leniusculus, 265, 957, 969, 970, 981,
983
nigrescens, 967
trowbridgii, 970
Paenecalyptonotus, 559, 645
Pagastia, 690
Palaemonetes, 961, 961, 962, 963, 964,
969, 983, 988
antrorum, 961
cummingi, 961, 969, 983
holthuisi, 961
intermedius, 969
kadiakensis, 956, 958, 959, 961, 962,
968
paludosus, 961, 962, 968
pugio, 969
Palaemonias, 961, 963, 964, 983, 985, 988
alabamae, 953, 961, 969, 974, 982
ganteri, 952, 961, 969, 974, 980, 982
paludosus, 957, 974
Palaeodipteron, 715
Paleochordodes protus, 280, 289
Palpomyia, 715
Paludicella, 509
articulata, 512, 517, 518
Panisopsis, 554, 578, 633, 640
setipes, 610
Panisus, 554, 633, 640
condensatus, 610
Pantala, 701
Paracamptus, 938
reductus, 950
reggiae, 950
Paracandona, 830, 833
euplectella, 810, 831
Paracapnia, 703
Parachordodes, 282, 292
lineatus, 291
violaceus, 291
Paracineta, 78
crenata, 79
Paractinolaimus, 274
Paracyatholaimus, 276
Paracyclops, 921, 936
canadensis, 949
chiltoni, 949
fimbriatus, 949
poppei, 949
smileyi, 949
yeatmani, 949
Paracymus, 684
Paradileptus, 85
robustus, 86
Paradiphascon, 529, 531
Paraeuglypha, 76
reticulata, 76
Paragnetina, 669
Paragonimus, 974
kellicoti, 974
westermani, 974
Paragordius, 292
varius, 282, 284
Paraleptophlebia, 662, 699, 739
Paralimnetis, 900
Paramastix, 51, 71
Paramecium, 43, 45, 48, 49, 51, 60, 62, 63,
89, 823
aurelia, 58, 59, 62
biaurelia, 58
bursaria, 62, 63
caudatum, 88
monoaurelia, 58
Paramideopsis, 558, 578, 635, 644
susanae, 601, 625, 630
Paranais, 437, 450
Paranyctiophylax, 672
Paraphaenocladius, 690
Paraphanolaimus, 265, 276
Paraphysomonas, 46, 67, 71
vestita, 66
Parascon, 529
Parastenocaris, 937
brevipes, 950
delamarei, 950
lacustris, 950
palmerae, 913
palmerae, 950
texana, 950
trichelata, 950
Parathyasella, 554, 640
heatherensis, 611
Parechiniscus, 542
Parhexapodibius, 529
Parophryoxus, 883
tabulatus, 884
Partnuniella, 578
Partuninia, 554, 640
steinmanni, 608
Pasithea, 859
Paulinella, 76
chromataphora, 76
Pectinatella, 509, 510
magnifica, 509 – 513, 520, 522
Pediastrum
duplex, 516
simplex, 516
Pedinomonas, 396
Pegias, 402, 406
fabula, 408
Pellioditis, 271, 273, 274
Pelocoris, 678, 709
Pelocypris, 829, 830, 835
alatabulbosa, 834
Pelomyxa, 67, 68, 71
palustris, 57, 75
Peltodytes, 684, 713
Peltoperla, 702, 740
Penardia, 75
granulosa, 75
Penardochlamys, 74
arcelloides, 75
Penelia, 852, 857, 873, 874
avirostris, 872
Penilia, 857, 873
Pentaqenia, 698
Taxonomic Index
Peranema, 62, 67, 71
trichophorum, 66
Perca flavescens, 366
Percymoorensis, 489, 496
kingi, 496
lateralis, 496
lateromaculata, 496
marmorata, 469, 480, 482, 496
septagon, 496
Pericantha, 859
truncata, 862
Peridinium, 66, 71
Perispira ovum, 63
Perlinella, 669
Perutilimermis, 278
Petalomonas, 71
abcissa, 66
Petrophila, 730
Phaenocoria, 158, 161, 162, 163, 170, 171
agassizi, 180
beauchampi, 159
falciodenticula, 180
highlandense, 180
kepneri, 180
lutheri, 180
typhlops, 158, 161
unipunctata, 158
virginiana, 180
Phagocata, 158, 168
velata, 168
Phagodrilus, 437, 439, 443
Phallodrilus hallae, 438, 454, 455
Pharyngostomoides procyonis, 459
Pheidole xerophila tucsonica, 2
Pherbella, 690
Pheromermis, 278
pachysoma, 260, 279
Philaster, 67
Philasterides, 87
armata, 86
Philobdella, 495
floridana, 495
gracilis, 495
Philodina, 197, 198, 218, 228, 235
acuticornis, 232
citrina, 217
Phoronis ovalis, 514
Phreatobrachypoda, 558, 646
multipora, 624
oregonensis, 624
Phryganella, 74
nidulus, 75
Phyllognathopus, 937
viguieri, 924, 950
Phylocentropus, 672, 705
Phymocythere, 830, 836
Physa, 300, 318, 975
vernalis, 306
Physella, 300, 304, 306, 307, 312, 318
anatina, 319
ancillaria, 304
cubensis, 302
gyrina, 306, 308, 319
heterostropha, 300
integra, 305, 319
vernalis, 301
virgata, 312, 319
zionis, 319
Physocypria, 830, 835
globula, 824
pustulosa, 815, 834
Piersigia, 553, 570, 579, 581, 586, 634, 639
limnophila, 607
Piguetiella, 451
Pilgramilla, 173
virginiensis, 179
Piona, 552, 557, 566, 568, 569, 572,
575 – 577, 579, 581, 582, 620, 638,
645, 648
carnea, 582, 597
constricta, 597
interrupta, 597
limnetica, 581
mitchelli, 597
napio, 574
Pionacercus, 557, 620, 645, 647
Pionopsis, 557, 638, 648
paludis, 619
Piscicola, 490, 494
geometra, 494
milneri, 494
punctata, 494
salmositica, 494
Piscicolaria, 468, 490
geometra, 481, 488, 489
milneri, 488, 489
punctata, 489
reducta, 490, 492, 494
salmositica, 489
Pisidium, 336, 343, 359, 360, 362, 368,
369, 370, 382, 383, 390, 399
amnicum, 348
annandalei, 369
casertanum, 348, 360, 369, 370, 383,
384, 389, 396
clarkeanum, 369
compressum, 345
conventus, 383
corneum, 389
crassum, 389
subtruncatum, 384
variabile, 345, 369
walkeri, 345, 346
Placobdella, 469, 488
costata, 481
hollensis, 468, 472, 488, 491
montifera, 488, 490, 491
mulilineata, 488, 492
nuchalis, 488, 490, 491
ornata, 472, 488, 491
papillifera, 474, 479, 486, 488, 492
parasticia, 472, 488, 491
pediculata, 474, 488, 491
phalera, 488
picta, 488
translucens, 488, 492
Plagiopyla, 89
nasuta, 88
Plagiopyxis, 74
callida, 75
1051
Planaria, 167
Planobella (Helisoma), 9
Planolineus, 174
Planorbella, 318
trivolvis, 313
Planorbis
contortus, 310
vortex, 306
Planorbula, 318
armigera, 304, 317
companulata, 319
trivolvis, 319
Platicrista, 529, 531, 532, 544
Platycentropus, 706
Platychirograpsus spectabilis, 960
Platycola, 81
longicollis, 80
Platyhydracarus, 559, 644
juliani, 603, 627
Plectocythere, 830, 836
Plectomerus, 402, 406
dombeyanus, 409, 413
Plectrocnemia conspersa, 824
Plectus, 256, 263, 265, 276
Pleocola, 538
Plethobasus, 402, 410
cyphyus, 406
Pleurobema, 402, 403, 411, 414
altum, 357
avellanum, 357
bourianum, 357
chattanoogaense, 357
clava, 406
collina, 367
flavidulum, 357
hagleri, 357
hanleyianum, 357
johannis, 357
murrayense, 357
nucleopsis, 357
rubellum, 357
troschelianum, 357
verum, 357
Pleurocera, 314, 323
acuta, 322
Pleuromonas, 71
jaculans, 72
Pleuronema, 87
coronatum, 86
Pleurotrocha, 245
Pleuroxus, 847, 859, 874, 878, 878
aduncus, 848
denticulatus, 856, 878
laevis, 869
procurvus, 856, 878, 879
trigonellus, 848
Plistophora aerospora, 228
Ploesma, 245, 247
Plumatella, 11, 510, 511, 512, 513, 514
bushnelli, 519, 521
casmiana, 508, 512, 513, 519, 521, 522
emarginata, 511, 512, 513, 519, 521
fruticosa, 511, 512, 513, 519, 520
fungosa, 511, 512, 513, 514, 522
nitens, 511, 512, 521
1052
Taxonomic Index
Plumatella, (continued)
nodulosa, 522
orbisperma, 512, 519
recluse, 511, 512
repens, 509, 511, 512, 513, 516, 519,
522
reticulata, 512, 519
rugosa, 522
simirepens, 522
vaihiriae, 511, 512, 513, 519, 521
Podiceps nigricollis, 39
Podon, 852, 857, 859, 887
leuckartii, 886
Podophyra, 79
fixa, 79
Podura, 717, 730
Polyarthra, 198, 201, 205, 208, 224, 227,
228, 245
dolichoptera, 223
major, 232
remata, 222
vulgaris, 223, 232
Polyartmiella, 890, 895
hazeni, 888
Polycelis, 161, 167
coronata, 168
tenuis, 161
Polycentropus, 13, 705
Polychaos timidum, 53
Polymerurus, 189, 190
Polymesoda, 397, 399
caroliniana, 399
Polypedilum, 690, 730
Polyphemus, 852, 857, 861, 862, 887
pediculus, 886
Polypodium, 6, 135, 149, 150, 151, 152
hydriforme, 140, 148 – 150
Polytoma, 71
Polytomella, 71
citri, 66
Pomacea, 9, 303, 323
paludosa, 324
Pomatiopsis, 320
lapidaria, 322
Pompholyx, 237, 239
Pontoporeia
affinis, 786, 789
femorata, 786
hoyi, 786
Popenaias, 402
popei, 406, 414
Porohalacarus, 648
Porolohmannella, 632, 648
Potamalpheops, 961
Potamilus, 363, 402, 410, 412, 414, 415
alatus, 406
purpuratus, 412
Potamocypris, 781, 815, 820, 822, 823,
830, 832
pallida, 816
smaragdina, 813, 815, 831
variegata, 819, 821
Potamogeton, 481, 986
Potamonemertes, 174
Potamopyrgus, 309
jenkinsi, 306
Potamothrix, 439, 453
moldaviensis, 439
Pottsiela erecta, 512, 517, 518
Prionchulus, 278
Prionocypris
canadensis, 829
glacialis, 829
Prismatolaimus, 274
Pristina, 439, 450
Pristinella, 439, 450
Proales, 202, 243
gigantea, 202
sordida, 217
werneckii, 243
Probopyrus pandalicola, 776
Probythinella, 320
Procambarus, 963, 964, 966, 967, 968,
972, 980, 982, 983, 988, 989
acutissimus, 987
acutus, 959, 977
clarkii, 972, 973, 975, 981, 984, 985
enoplosternum, 957
fallax, 978
gracilis, 978, 980, 987
hagenianus, 973
lunzi, 953, 957
paeninsulanus, 959, 978, 986
pecki, 987
primaevus, 964
pubescens, 972
pygmaeus, 985
simulans, 957
spiculifer, 973
tenuis, 960
versutus, 957
zonangulus, 972, 973
Procaris, 961
Procladius, 734, 735
Procotyla, 169
fluviatilis, 168
Prodesmodora, 265, 276
Progomphus, 666
Proichthydium, 185, 190
Proichthyoides, 185
Promenetus, 307, 318
exacuous, 304, 319
Prorhynchella, 169
minuta, 171, 180
Prorhynchus, 157, 161, 164, 166
stagnalis, 161, 179
Prorodon, 67, 85
teres, 86
Prosimulium, 690, 730
Prosotoma, 6
Prostoma, 7, 172, 174, 175, 176,
265
asensoriatum, 175
canadiensis, 174, 175, 176
eilhardi, 174, 176
graecense, 174, 176
rubrum, 175
Protilimnesia, 556, 617, 643
Protoascus, 170
wisconsinensis, 180
Protocaris, 867
Protoneura, 701
Protoplasa, 716
Protzia, 554, 565, 571, 578, 608, 633, 640
Psammonitocrella, 937
boultoni, 950
longifurcata, 950
Psammoryctides, 453, 454
barbatus, 439
Psephenus, 687, 713, 730
Pseudechiniscus, 529, 530, 538, 542
Pseudiron, 699
Pseudobiotus, 529, 531, 532, 533, 538,
539, 543
augusti, 536
kathmanae, 536
macronyx, 537
megalonyx, 536, 537
Pseudocandona, 822, 830, 833
albicans, 822
Pseudocentroptiloides, 662
Pseudochordodes, 292
Pseudochydorus, 874, 878
globosus, 862, 879
Pseudodiaptomus, 934, 934
Pseudodifflugia, 75
gracilis, 75
Pseudodiphascon, 529, 531, 532, 533, 534,
543
Pseudofeltria, 557, 638, 645, 647
multpora, 596, 620
Pseudohexapodibius, 529, 531, 532
Pseudohydryphantes, 564, 579, 639
Pseudomermis, 278
Pseudomicrothorax, 67, 89
agilis, 80
Pseudomoina, 883
lemnae, 885
Pseudophaenocora, 170
sulflophila, 180
Pseudoploesma, 245, 247
Pseudoprorodon, 85
ellipticus, 86
Pseudosida, 874
bidentata, 872
Pseudosuccinea, 320
collumella, 304, 306, 310, 321
Pseudotorrenticola, 555
chiricahua, 614
Psilotreta, 672, 675, 706
Psilotricha, 83
acuminata, 82
Psychoda, 715
Psychomyia, 672, 705
Pternarcys, 730
Pterodrilus, 460
alcicornis, 460
Ptychobranchus, 402, 406, 409, 412, 415
Ptychoptera, 716
Ptygura, 202, 214, 223, 224, 237
barbata, 238
beauchampi, 205, 214
Taxonomic Index
mucicola, 238
Punctodora, 276
Pycnopshche, 672
Pyganodon, 363, 402, 403, 408, 412
cataracta, 367, 376, 382, 384
grandis, 348 – 352, 358 – 361, 366, 373,
375, 386, 407
Pyrgulopsis, 303, 320
Pyxicola, 81
affinis, 80
Pyxidicula, 74
operculata, 75
Quadrula, 9, 363, 402, 410, 413, 414
cylindrica, 410
quadrula, 407
tuberosa, 357
Quadrulella, 74
symmetrica, 75
Quincuncina, 402, 407, 413
Quistadrilus, 439, 452
Radiospongilla, 117, 130
cerebellata, 124, 125, 130
crateriformis, 124, 125, 130
Radix, 320
auricularia, 321
Ramajendas, 529, 531, 532, 538, 539
Ramazzottius, 529, 531, 532, 537,
543
Rana
clamaitans, 825
temporaria, 288
Rangia cuneata, 385, 386, 397
Raphidiophrys, 78
elegans, 77
Rectocephala, 169
Redudasys, 185
fornerise, 182
Regina alleni, 973
Reticulomyxa, 77
filosa, 76
Rhabditolaimus, 274
Rhabdolaimus, 278
brachyuris, 263
Rhabdostyla, 81
pyriformis, 80
Rhadinocythere, 830, 837
Rhagovelia, 678
Rhantus, 684
Rhapinema, 320
Rheocyclops, 937
carolinianus, 949
hatchiensis, 949
talladega, 949
virginianus, 949
Rheumatobates, 678
Rhinoglena, 215, 224, 243
fertoensis, 221
Rhithropanopeus harrisii, 960
Rhizodrilus, 436, 439
lacteus, 453
Rhodacmea, 316
rhodacme, 317
Rhyacodrilus, 439, 454
brevidentatus, 454
falciformis, 438, 455
sodalis, 440
Rhyacophila, 706, 730
Rhynchelmis, 448
brooksi, 448, 449
Rhyncholimnochares, 553, 563, 569, 570,
574, 578, 579, 607, 634, 639
kittatinniana, 587
Rhynchomesostoma, 162, 170
rostratum, 171, 180
Rhynchomonas, 71
nasuta, 72
Rhynchoscolex, 157, 158, 162, 165,
172
pegephilum, 159
platypus, 179
simplex, 165, 179
tauricum, 179
Rhynchotalona, 874, 876
falcata, 875
Richtersius, 529, 531, 532
Ripistes, 438, 451
parasita, 450
Romanomermis, 280
culicivorax, 269
Rotaria, 198, 235
Rutripalpus, 555, 612, 641
Saccamoeba, 74
lucens, 73
Safilippodytes, 684
Sagittocythere, 830, 837
Salda, 709
Salmasellus steganothrix, 792
Salmo trutta, 474
Salpingoeca, 71
fusiformis, 72
Salvelinus alpinus, 824
Saprodinium, 83
dentatum, 82
Sarscypridopsis, 830, 832
Sarsilatona, 874
serricaudata, 872
Sathodrilus, 460
attenuatus, 461
carolinensis, 461
hortoni, 461
Saurocythere, 830, 836
Scapholeberis, 34, 143, 850, 859, 881
kingi, 226
rammneri, 880
Scapulicambarus, 967
Scelorchilus rebecula, 290
Scenedesmus, 225, 396, 870, 931
costato-granulatus, 229
Sciomyza, 690
Scottia, 830, 833
Scutolebertia, 555, 642
trinitensis, 615
1053
Scyphidia, 81
physarum, 80
Scyphozoa, 145
Seinura, 270
Seison, 233
Selenastrum, 396, 870
Semigordionus, 292
Senecella, 917, 934
calanoides, 923, 948
Sepedon, 690, 715
Serratella, 662
Setnonema, 699
Setodes, 672
Setopus, 191
dubius, 187
Setvena, 669
Sialis, 707, 730
fuliginosa, 824
Sida, 850, 851, 852, 857, 859, 861, 873
crystallina, 872
Simocephalus, 143, 847, 850, 859, 881
serrulatus, 880
vetulus, 880
Simposonaias, 402
ambigua, 363, 407, 411
Simulium, 26, 690, 716, 730, 740, 760
Sinantherina, 8, 211, 212, 215, 224, 227,
237
socialis, 212, 214, 216, 217, 228,
232
spinosa, 227
Sineportella forbesi, 512, 518
Sinobosmina, 881, 882
fatalis, 882
Sinocalanus, 935
doerri, 948
Sinodiaptomus, 935
sarsi, 948
Siolineus, 174
Siphlonurus, 699
Siphloplecton, 699
Siphonophanes grubei, 892
Siqara, 709
Siskiyouthyas, 554, 640
Sisyra, 730
Skistodiaptomus, 935
bogalusensis, 948
carolinensis, 948
mississippiensis, 948
oregonensis, 948
pallidus, 948
pygmaeus, 948
reighardi, 948
sinuatus, 948
Slavina, 452
appendiculata, 451
Sminthurides, 730
Soldanellonyx, 648
Solenopsis invicta, 373, 385
Soliperda, 669
Somatochlora, 666
Somatogyrus, 320
Somersiella, 961
Sparganophilus, 446
1054
Taxonomic Index
Spathidium, 85
spathula, 84
Specaria, 452
Spelaedrilus, 449
multiporus, 449
Sperchon, 552, 555, 572, 579, 612, 635,
642
glandulosus, 588
Sperchonopsis, 555, 635, 642
ecphyma, 561, 562
nova, 612
Sphaerium, 343, 359, 360, 362, 368, 383,
390, 400
corneum, 348, 373, 400
fabule, 369, 400
nitidum, 400
occidentale, 350, 351, 400
simile, 348, 400
striatinum, 345, 346, 347, 369, 381, 383,
388, 389, 400
Sphaeroeca, 71
volvox, 72
Sphaerophyra, 79
magna, 79
Sphagnum, 241, 244
Sphalloplana, 167
Sphenoderia, 76
lenta, 76
Spilochlamys, 320
Spinalona, 847
Spirosperma, 436, 452, 453
Spirostomum, 55, 85
minus, 84
Spirulina, 827
Spongilla, 129
alba, 106, 107, 125, 129
aspinosa, 125, 129
cenota, 125, 126, 129
lacustris, 98, 99, 103 – 114, 117, 126,
128, 129
Spumella, 66
Squalorophyra, 81
macrostyla, 79
Squatinella, 241
Stachyamoeba, 73
lipophora, 73
Stagnicola, 320
catascopium, 314
elodes, 314, 321
emarginata, 314
palustris, 314
Stenelmis, 13, 684, 712, 713
Stenochironomus, 690
Stenocypria longicomosa, 829
Stenocypris, 830, 833
major, 825
Stenonema, 662
Stenoperla prasina, 289
Stenophylax, 288
Stenophysa, 318, 319
Stenostomum, 157, 159, 161, 163, 165
anatirostrum, 179
anops, 179
arevaloi, 179
beauchampi, 166, 179
brevipharyngium, 166, 179
ciliatum, 179
cryptops, 179
glandulosum, 166, 179
grande, 179
kepnere, 179
leucops, 158, 162, 166, 179
mandibulatum, 179
membranosum, 179
occultum, 179
pegiphilum, 179
predatorium, 161, 179
pseudoacetabulum, 179
simplex, 179
temporaneum, 179
tuberculosum, 179
unicolor, 179
uronephrium, 179
ventronephrium, 179
virginianum, 179
Stentor, 5, 55, 65, 83
polymorphous, 84
Stephanella, 514
hina, 511, 518, 519
Stephanoceros, 214, 224, 235
Stephanodiscus, 974
Stephensoniana, 450
Stichotricha, 83
aculeata, 82
Stictotarsus, 684
Stokesia, 87
vernalis, 88
Stolonicyclops, 936
heggiensis, 949
Strandesia, 830, 833
canadensis, 822, 829
Stratospongilla penneyi, 126, 129
Streblocerus, 883
serricaudatus, 884
Strelkovimermis, 280
Streptocephalus, 891, 892, 894, 897
dichotomus, 893
mackini, 891, 893
seali, 890, 891
texanus, 888
Striobia, 320
Strobilidium, 62
Strombidinopsis, 83
setigera, 82
Strombidium, 83
viride, 82
Strombidum, 63
capitatum, 60
Strombidium, 83
gyrans, 82
Strongylidium, 82
crassum, 82
Strongylostoma, 170, 172
elongatum, 180
radiatum, 180
simplex, 171
Strophitus, 402, 403, 412
undulatus, 407, 408
Strophopteryx, 703
Stygalbiella, 558, 646
yavapai, 621
Stygameracarus, 559, 644
cooki, 627
Stygobromus, 786, 787, 792
canadensis, 792
Stygohalacarus, 648
Stygomomonia, 558, 635, 643
mitchelli, 601, 625
riparia, 625
Stygonitocrella, 937
sequoyahi, 950
Stygothrombium, 553, 585, 588, 606, 634,
638
Stylaria, 438
lacustris, 450, 451
Stylochaeta, 7, 191
Stylocometes, 81
digitalis, 79
Stylodrilus, 443, 449
californianus, 449
heringianus, 440, 441, 449
sovaliki, 450
Stylonyhcia, 81
mytilus, 82
Styloscolex, 449
opisthothecus, 450
Styraconyx hallasi, 529, 538
Submiraxona, 558, 647
gilana, 622
Suomina, 165
turgida, 165
Suphisellus, 684
Suragina, 693
Sympetrium, 666
Syncaris, 963, 964, 983, 985, 988
pacifica, 961, 969, 983
pasadenae, 961
Synchaeta, 195, 198, 201, 202, 210, 223,
227, 228, 245, 247, 928
cecelia, 232
oblongata, 215, 222, 226
pectinata, 209, 215, 222, 228
Synedra, 974
Synendotendipes, 690
Tachidus, 937
Tachopteryx, 701
Taeniopteryx, 669, 703
Taeniorhynchus, 732
Tanytarsus, 730, 735
Taphromysis louisiannae, 786, 788
Tartarothyas, 554, 564, 571, 578, 611, 633,
641
Tasserkidrilus, 454
Tegeocranellus, 649
Telmatodrilus, 452, 455
onegensis, 452
vejdovskyi, 436, 454
Temora, 923
Tenagodrilus musculus, 450
Tendipes pectinatellae, 513
Taxonomic Index
Teneridrilus mastix, 454, 455
Teratocephalus, 276
Teretifrons, 874
Testudacarus, 555, 591, 636, 641
americanus, 614
Testudinella, 237, 239
Tetanocera, 690
Tetrahymena, 43, 45, 48, 51, 57, 60, 61,
62, 65, 67, 89
pyriformis, 45, 58, 88
Tetrakentron, 538
Tetramermis, 278
Tettodrilus, 460
Teutonia, 552, 555, 569, 579, 612, 635,
641
lunata, 589
Thamnocephalus, 895
platyrus, 888
Thaumalea, 690, 716
Thecacineta, 78
cothurniodes, 79
Thecamoeba, 53, 67, 74
sphaeronucleolus, 74
Theodoxus, 307
Theristus, 265
pertenuis, 263
Thermacarus, 555, 578, 633, 641
nevadensis, 565, 611
Thermastrocythere, 830, 837
Thermobatynella adami, 781
Thermocyclops, 929, 936
crassus, 949
inversus, 949
parvus, 949
tenuis, 949
Thermonectus, 684
Thermosbaena mirabilis, 781
Thermosphaeroma
subequalum, 781
thermophilum, 781, 792
Thermozodium, 529
esakii, 529, 539
Theromyzon, 474, 488, 493
biannulatum, 488
bifarium, 488, 494
maculosum, 488, 493
rude, 475, 488, 494
sexoculatum, 488
tessulatum, 477, 481, 488, 493
trizonare, 474, 477, 478, 479, 481, 486,
488, 492, 493
Thiara, 323
granifera, 322
Thinodrilus, 448
Thraulodes, 662
Thulinia, 529, 531, 532, 533, 539, 541, 543
Thuricola, 81
folliculata, 80
Thyas, 554, 560, 569, 572, 578, 579, 581,
633, 641
barbigera, 569
rivalis, 611
stolli, 565, 611
Thyasella, 554, 640
Thyasides, 554, 611, 633, 641
Thyopsella, 554, 640
dictyophora, 610
occidentalis, 610
Thyopsis, 554, 640
cancellata, 610
Thyopsoides, 554, 640
plana, 610
Tiguassu, 442
Tillina, 89
magna, 88
Timpanoga, 662
Tintinnidium, 83
fluviatile, 82
Tintinnopsis, 83
cylindricum, 82
Tiphys, 557, 576, 577, 579, 619, 638,
648
americanus, 574, 596
ornatus, 596
Tipula, 730, 740, 760
Tobrilus, 265, 275, 278
gracilis, 263
grandipapellatus, 263
Tokophyra, 81
quadripartita, 63, 64, 79
Tonnacypris, 823, 830, 833
glacialis, 821, 829
Torrenticola, 552, 555, 572, 579, 592, 614,
636, 641
Toxolasma, 402, 408
parvus, 351, 375
texasiensis, 348, 353, 382, 407
Trachelius, 85
ovum, 86
Trachelophyllum, 85
apiculatum, 84
Trajancypris, 828, 830, 833
Tramea, 701
Traverella, 662
Trepobates, 678
Trepomonas, 57
Triannulata, 460
magna, 460
Trianodes, 672
Tribelos, 690
Trichamoeba, 74
cloaca, 73
Trichocera, 244, 246
pusilla, 205, 217
rattus, 202, 224
Trichocorixa, 678, 730
Trichodina, 81
pediculis, 80
Trichodrilus, 443, 449
allegheniensis, 448
Trichogramma, 730
Trichophyra, 79
epistylidis, 79
Trichothyas, 554, 640
muscicola, 609
Trichotria, 243, 244
Tricorythodes, 698
Trilobus, 265
1055
Trinema, 76
enchelys, 76
Triops, 15, 889, 890, 891, 892, 894, 900
longicaudatus, 899
Tripyla, 265, 278
Tritogonia, 402, 407
verrucosa, 409, 413
Troaculus, 229
Trochodrilus, 438
Trochosphaera, 237, 239
Trochospongilla, 120
horrida, 106, 126, 127, 130
leidii, 127, 129
pennsylvanica, 106, 108, 127, 130
Troglocambarus, 964, 966, 967, 968, 988
maclanei, 966, 976
Troglocubanus, 961
Tropisternus, 684, 711, 712, 713
Tropocyclops, 227, 922, 936
extensus, 949
jerseyensis, 949
prasinus, 917, 924, 927, 949
Truncilla, 402, 411, 414
truncata, 343, 407
Trypanosoma, 52
Tubifex, 163, 439, 454
tubifex, 437, 438, 439, 445, 455
Tubificoides benedii, 437, 439
Tulotoma, 303, 323
magnifica, 324
Turaniella, 89
vitrea, 88
Turbanella hyalina, 187
Tvetenia, 690
Tylenchus, 270
Tylocephalus, 263
Typha, 816
Typhlatya, 961
Typhloplana, 162, 164, 170
viridata, 158, 160
Typhloplanella, 170
halleziana, 180
Tyrrellia, 556, 593, 617, 636, 643
Uchidastygacarus, 559, 579, 643
acadiensis, 626, 630
ovalis, 626
Uglukodrilus, 460
Uncinais uncinata, 450
Uncinocythere, 830, 837
Undula, 185, 189, 190
Unio, 363
pictorum, 341, 367, 382
tumidus, 348, 367, 376, 391
Uniomerus, 402, 412, 415
caroliniana, 408
tetralasmus, 351, 373, 375, 407, 412
Unionicola, 384, 556, 567 – 569, 574, 577,
579, 581, 582, 594, 618, 637, 648
crassipes, 575
intermedia, 577
Urceolaria, 81
mitra, 80
1056
Taxonomic Index
Urceolus, 71
cyclostomas, 66
Urnatella gracilis, 11, 515, 516, 517
Urocentrum, 87
turbo, 88
Uroglena, 71
americana, 72
Uroleptus, 81
piscis, 82
Uronema, 87
griseolum, 86
Urostyla, 81
grandis, 82
Urotricha, 85
farcta, 84
Urozona, 87
bütschlii, 86
Utaxatax, 555, 569, 634, 642
newelli, 589
ovalis, 613
Utricularia, 231
Utterbackia, 402, 403, 409, 413
imbecillis, 384, 407
Vaginicola, 81
ingenita, 80
Valkampfia, 73
avaria, 73
Valvata, 309, 320
sincera, 321
tricarinata, 310, 321
Vampyrella, 75
lateritia, 76
Vanella miroides, 74
Varichaetadrilus, 453, 454, 454
Varsoviella kozminski, 159
Vasicola, 85
ciliata, 84
Vaucheria, 243
Vejdovskyella, 438, 451, 452
Velesunio ambiguus, 348
Venustaconcha, 402
ellipsiformis, 407, 412
Vexillifera telemathalassa, 73
Vibrio cholerae, 929
Victorella pavida, 512, 518
Villosa, 364, 402, 408, 411, 412,
414, 415
constricta, 408
iris, 382, 383
villosa, 407
Viviparus, 303, 306, 323
georgianus, 305, 306, 324
subpurpureus, 309
Volsellacarus, 559, 636, 644
sabulonus, 626, 630
Volvox, 244
Vorticella, 5, 55, 64, 81
campanula, 80
Vorticifex, 318
effusa, 319
Wandesia, 554, 560, 565, 569, 570, 571,
578, 579, 608, 633, 641
thermalis, 569
Wettina, 557, 579, 637, 647
octopora, 619
ontario, 595
Wlassicsia, 883
kinistenensis, 885
Woolastookia, 558, 637, 646
pilositarsa, 599, 623
setosipes, 599
Xerobiotus, 529, 531, 532, 537
pseudohufelandi, 537
Xiphocentron, 705
Xironodrilus, 460
Xironogiton, 459
instabilis, 460
victoriensis, 458
Yachatsia, 559, 645
mideopsoides, 626
Yagerocaris, 961
Ylodes, 672
Zoothamnium, 59, 81
arbuscula, 80
Zschokkea, 554, 611, 633, 641