ORIGINAL RESEARCH ARTICLE
published: 30 November 2012
doi: 10.3389/fmicb.2012.00402
Microenvironmental ecology of the chlorophyll
b-containing symbiotic cyanobacterium Prochloron in the
didemnid ascidian Lissoclinum patella
Michael Kühl 1,2,3 *, Lars Behrendt 1 , Erik Trampe 1 , Klaus Qvortrup 4 , Ulrich Schreiber 5 , Sergey M. Borisov 6 ,
Ingo Klimant 6 and Anthony W. D. Larkum 2
1
2
3
4
5
6
Marine Biological Section, Department of Biology, University of Copenhagen, Helsingør, Denmark
Plant Functional Biology and Climate Change Cluster, University of Technology Sydney, Sydney, NSW, Australia
Singapore Centre on Environmental Life Sciences Engineering, School of Biological Sciences, Nanyang Technological University, Singapore
Department of Biomedical Sciences, Core Facility for Integrated Microscopy, University of Copenhagen, Copenhagen, Denmark
Julius-von-Sachs Institut für Biowissenschaften, Universität Würzburg, Würzburg, Germany
Department of Analytical and Food Chemistry, Technical University of Graz, Graz, Austria
Edited by:
Hans-Peter Grossart,
IGB-Leibniz-Institute of Freshwater
Ecology and Inland Fisheries,
Germany
Reviewed by:
Heather Bouman, University of
Oxford, UK
Rebecca J. Case, University of
Alberta, Canada
*Correspondence:
Michael Kühl , Marine Biological
Section, Department of Biology,
University of Copenhagen,
Strandpromenaden 5, DK-3000
Helsingør, Denmark.
e-mail: mkuhl@bio.ku.dk
The discovery of the cyanobacterium Prochloron was the first finding of a bacterial oxyphototroph with chlorophyll (Chl) b, in addition to Chl a. It was first described as Prochloron
didemni but a number of clades have since been described. Prochloron is a conspicuously large (7–25 µm) unicellular cyanobacterium living in a symbiotic relationship, primarily
with (sub-) tropical didemnid ascidians; it has resisted numerous cultivation attempts and
appears truly obligatory symbiotic. Recently, a Prochloron draft genome was published,
revealing no lack of metabolic genes that could explain the apparent inability to reproduce
and sustain photosynthesis in a free-living stage. Possibly, the unsuccessful cultivation
is partly due to a lack of knowledge about the microenvironmental conditions and ecophysiology of Prochloron in its natural habitat. We used microsensors, variable chlorophyll
fluorescence imaging and imaging of O2 and pH to obtain a detailed insight to the microenvironmental ecology and photobiology of Prochloron in hospite in the didemnid ascidian
Lissoclinum patella. The microenvironment within ascidians is characterized by steep gradients of light and chemical parameters that change rapidly with varying irradiances. The
interior zone of the ascidians harboring Prochloron thus became anoxic and acidic within a
few minutes of darkness, while the same zone exhibited O2 super-saturation and strongly
alkaline pH after a few minutes of illumination. Photosynthesis showed lack of photoinhibition even at high irradiances equivalent to full sunlight, and photosynthesis recovered
rapidly after periods of anoxia. We discuss these new insights on the ecological niche of
Prochloron and possible interactions with its host and other microbes in light of its recently
published genome and a recent study of the overall microbial diversity and metagenome
of L. patella.
Keywords: Prochloron, symbiosis, microenvironment, photobiology, microsensor, bioimaging, didemnid ascidian,
Lissoclinum patella
INTRODUCTION
The phylum Cyanobacteria harbors a diversity of morphotypes
ranging from minute <1 µm unicellular forms (Prochlorococcus spp.) up to 100 µm thick filaments (Oscillatoria spp.) and
large colonial cell aggregations visible by the naked eye (Palinska, 2008). Cyanobacteria are the only oxyphototrophs among
prokaryotes and are considered inventors of oxygenic photosynthesis >2.5 billion years ago, major drivers in the formation of
the first biological communities in the fossil record, i.e., microbial mats and stromatolites, as well as key architects of the present
biosphere through their photosynthetic O2 production leading
to an oxic atmosphere over geologic time scales. Cyanobacteria
are also the only oxyphototrophs capable of N2 fixation, either
in specialized cells (heterocysts) or via spatio-temporal modulation of their metabolic activity (Stal and Zehr, 2008), a trait
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that is employed by several protists, plants, and animals harboring symbiotic cyanobacteria (Adams, 2000; Lesser et al., 2004). In
recent years, the analysis of cyanobacterial genomes has revealed
a large degree of genetic exchange and plasticity inside this phylum (Zhaxybayeva et al., 2006), and cyanobacteria are regarded
as prime candidates involved in the endosymbiosis leading to the
evolution of algae and higher plants (Price et al., 2012).
Cyanobacteria also harbor species exhibiting unique characteristics very different from “typical” cyanobacteria. Most notably,
this includes the prochlorophytes and the Chl d-containing
cyanobacteria in the genus Acaryochloris that exhibit fundamental
differences in terms of pigmentation, structure, and properties of
their photosynthetic apparatus (Partensky and Garczarek, 2003;
Larkum and Kühl, 2005; Kühl et al., 2007a). More recently, findings of the new chlorophyll f in cyanobacterial enrichments from
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Kühl et al.
Prochloron microenvironment
stromatolites (Chen et al., 2010), and a widespread but yet uncultivated diazotrophic cyanobacterium without a functional PSII
(Bothe et al., 2010) add on to the concept of cyanobacteria being
a genetic melting pot and origin for a wide variety of photosynthetic adaptations (Larsson et al., 2011; Schliep et al., 2012). The
ecology of such exotic cyanobacteria is not well understood, and
in this study we focus on the ecology and habitat characteristics of
the conspicuous symbiotic cyanobacterium Prochloron spp., which
can be found and harvested in large quantities from their tunicate
hosts (so-called didemnid ascidians) on coral reefs and mangrove
systems but has resisted all cultivation attempts since its discovery
in 1975 (Lewin and Cheng, 1989).
The taxonomy of Prochloron remains ambiguous. The type
species was initially called Synechocystis didemni and later renamed
Prochloron didemni (Lewin, 1977). Detailed comparative electron
microscopy of different symbiont-host associations indicated several characteristic morphotypes hypothesized to represent different phylotypes of Prochloron (Cox, 1986). However, molecular data
comparing sequences of the Prochloron 16S rRNA gene obtained
from different didemnid ascidians and geographic location indicate a fairly strong global similarity (Münchhoff et al., 2007) and a
genomic study of Prochloron samples along a >5000 km transect in
A
the Pacific showed >97% identity and strong synteny of Prochloron
genomes (Donia et al., 2011a,b).
Cyanobacteria in the genus Prochloron are ∼7–25 µm wide
(Cox, 1986), bright green spherical cells with stacked thylakoids
oppressed to the cell periphery (Figure 1); they were discovered
in close association with (sub)tropical didemnid ascidians (Lewin
and Cheng, 1975; Newcomb and Pugh, 1975). While the presence of microbial phototrophs and O2 production in didemnid
ascidians was well known (Maurice, 1888; Smith, 1935; Tokioka,
1942), it was a surprise that Prochloron contained no phycobilins
but Chl b in addition to Chl a (Lewin and Withers, 1975). At
that time, Chl b was only known from algae and higher plants,
and Prochloron was for some time regarded a missing link in
chloroplast evolution. However, molecular phylogenetic studies
have unequivocally shown that Prochloron is but one of several
lineages in the cyanobacteria, wherein Chl b has evolved (Palenik
and Haselkorn, 1992; La Roche et al., 1996).
Along with the other two groups of prochlorophytes, i.e.,
Prochlorococcus and Prochlorothrix, Prochloron shares several
additional differences to other cyanobacteria, including special
membrane-bound Chl a/b light harvesting complexes that are
unrelated to the Chl a/b antenna in the light harvesting complex
B
Branchial aperture
Cloacal aperture
Cloacal cavity
Tunic
Zooid
PROCHLORON
Biofilm
C
E
D
5 µm
t
t
s
m
s
s
20 µm
FIGURE 1 | The Prochloron -Lissoclinum patella symbiotic
association. (A) A colony (∼20 cm2 ) of the didemnid ascidian
Lissoclinum patella covering corals with its 5–10 mm thick opaque
cartilaginous tunic, i.e., a matrix of protein and cellulose-like
carbohydrates containing calcareous spicules. (B) A schematic drawing
of a cross-section illustrating the organization of zoids and symbionts in
the tunic. The zooids are embedded in the tunic, where they suck in
and filtrate particles out of the seawater. Waste products and filtered
Frontiers in Microbiology | Aquatic Microbiology
200 µm
water are excreted into the surrounding peribranchial space and cloacal
cavities in the colony tunic and eventually expelled via a joint cloacal
aperture. (C) Individual Prochloron cells extracted from the ascidians,
note the peripheral arrangement of thylakoids in the cells. (D,E)
Electron micrographs (D) [TEM, (E) FIB SEM] of the symbiont-host
interface in L. patella showing round Prochloron cells (s) embedded
into the host tunic (t) and a fibrous exopolymeric substance (m, arrow).
(C) Was redrawn and modified after Maruyama et al. (2003).
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Kühl et al.
of eukaryotic oxyphototrophs (La Roche et al., 1996; Partensky
and Garczarek, 2003). Interestingly, genes encoding for the socalled “prochlorophyte Chl b-binding” proteins (Pcbs) are found
both in prochlorophytes and in the Chl d-containing cyanobacterium Acaryochloris marina, which is not closely related to the
prochlorophytes on basis of their 16S rRNA gene phylogeny, indicating a more ancient origin of these genes or lateral gene transfer
(Chen et al., 2005).
Prochloron forms symbiotic associations with didemnid ascidians in the genera Didemnum, Trididemnum, Lissoclinum, and
Diplosoma and represents the only known obligate photosymbiosis in the phylum Chordata (Hirose et al., 2009a). Studies of
the 18S rRNA-based molecular phylogeny of didemnid ascidians
indicate that photosymbiosis with Prochloron has occurred independently in each of these genera (Yokobori et al., 2006). Besides
the unsuccessful attempts to cultivate Prochloron separated from
its hosts, a symbiotic relation is supported, e.g., by observations
of carbon and nitrogen exchange between Prochloron and host
(Lewin and Pardy, 1981; Kremer et al., 1982; Griffiths and Thinh,
1983; Koike et al., 1993), light-enhanced growth of didemnids
with Prochloron (Olson, 1986), and fascinating specializations in
didemnids ensuring vertical transmission of Prochloron to ascidian larvae during maturation and before they leave the parent
tunic (Hirose, 2009), i.e., the extracellular matrix of protein and
cellulose-like carbohydrates, wherein the host zoids are embedded.
However, the exact nature of the symbiosis and mutual benefits
in the ascidian-Prochloron association is still poorly understood
(Hirose and Maruyama, 2004).
Different modes of association between Prochloron and didemnid ascidians have been described (Cox, 1986; Hirose et al.,
2009a) including (i) colonization of the outer surface and upper
tunic, (ii) colonization of the inner cloacal cavities and the peribranchial space of zoids, (iii) a more wide spread colonization
in the tunic, and (iv) intracellular Prochloron in some didemnid
species. Additionally, there is macro- and microscopic evidence
that Prochloron can be associated with the surface of holothurians (Cheng and Lewin, 1984), sponges (Parry, 1986), and a range
of non-didemnid ascidians (Cox, 1986). Significant amounts of
suspended Prochloron in seawater have been observed (Cox, 1986)
and molecular surveys indicated that Prochloron could be thriving in stromatolites (Burns et al., 2004), but confirmation of such
potentially free-living Prochloron awaits further direct evidence of
their actual presence and ecological niche in such systems.
Detailed physiological and photobiological studies of
Prochloron are few (reviewed in Kühl and Larkum, 2002) and have
mostly involved measurements on solute exchange or photosynthetic performance of intact ascidian-Prochloron associations or
on extracted Prochloron cells, which only remain photosynthetically competent for a few hours to a day (Critchley and Andrews,
1984). This has hampered both cultivation attempts and a more
detailed understanding of the ecological niche of Prochloron in hospite. However, some niche characteristics are well known such as
the presence of UV absorbing mycosporin-like compounds that
act as sunscreens (Dionisio-Sese et al., 1997; Maruyama et al.,
2003; Hirose et al., 2006), the presence of enzymes for detoxification of reactive oxygen species (ROS; Lesser and Stochaij, 1990),
and the prevalence of a diversity of highly bioactive secondary
metabolites (especially cyanobactins) in Prochloron that may, e.g.,
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Prochloron microenvironment
have allelopathic functions against other phototrophs (Schmidt
and Donia, 2010; Donia et al., 2011b; Schmidt et al., 2012).
The photosynthetic apparatus of Prochloron appears similar
to other cyanobacteria with conventional electron transport reactions through photosystem (PS) I and II and a typical oxygen
evolution complex. However, special light harvesting supercomplexes based on the prochlorophyte chlorophyll a/b (pcb) protein
are present in both PSI and PSII (Bibby et al., 2003; Murray et al.,
2006). These light harvesting systems give rise to a highly efficient
photosynthetic electron transport system, where the maximum
quantum yield of PSII approaches 0.82, similar to that in higher
plants and much higher than in most cyanobacteria (Schreiber
et al., 1997, 2002).
Carboxysomes are present, and a significant part of the carbonic anhydrase present is found in these bodies (Griffiths, 2006)
along with all the Rubisco enzyme (Swift and Leser, 1989), which
is responsible for fixing CO2 to phosphoglyceric acid. There
appears to be an active supply of organic products to the ascidian host and these appear to be early carbon products of the
Calvin–Benson cycle (Kremer et al., 1982). This translocation is
host-dependent and represents about 12–56% of reduced carbon for host respiration (Olson and Porter, 1985; Alberte et al.,
1987).
Other aspects of metabolism, e.g., the source and dynamics of
N2 fixation and nitrogen turnover in the ascidian-Prochloron symbiosis are still unresolved and debated (e.g., Odintsov, 1991; Donia
et al., 2011b). There is evidence that in addition to carbon, nitrogen
is also recycled in the symbiosis (Koike et al., 1993). Ammonium
is the major nitrogenous waste of the host (Goodbody, 1974) and
is taken up by Prochloron (Parry, 1985). It has been proposed that
N2 fixation contributes to the nitrogen requirements (Paerl, 1984).
This was disputed by Parry (1985), but is supported by nitrogen
isotope ratios in host and Prochloron, which are consistent with
nitrogenase activity (Kline and Lewin, 1999).
The recent publications of the first draft Prochloron genome as
well as studies of the microbiome of photosymbiotic didemnids
(Donia et al., 2011a,b; Behrendt et al., 2012a) have now added significant new insights to the ecology of Prochloron, and potential
interactions with its didemnid hosts and other members of the
microbiome. However, the interpretation of such molecular surveys in terms of physiology and metabolic interactions mostly rely
on genomic evidence that still needs experimental verification for
Prochloron under natural conditions.
While many studies have speculated about the microenvironmental conditions of Prochloron in hospite, very few experimental
data on the microenvironment in didemnid ascidians have been
published (Kühl and Larkum, 2002; Behrendt et al., 2012a). These
data indicate highly dynamic physico-chemical conditions that
are strongly modulated by irradiance. In this study, we present
new data and review the current knowledge about the microenvironment and metabolic activity of symbiotic Prochloron associated with didemnid ascidians. We focus on the large colonial
didemnid ascidian Lissoclinum patella, which covers significant
areas of substratum on coral reefs with an opaque cm-thick
cartilaginous tunic harboring large quantities of Prochloron in
the internal peribranchial and cloacal cavities of the filter-feeding
zoids (Figure 1). The tunic of L. patella is an integumentary extracellular matrix of protein and cellulose-like carbohydrates covered
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Kühl et al.
by a dense tunic surface cuticle containing calcareous spicules as
well as a diversity of specialized tunic host cells (Goodbody, 1974;
Hirose, 2009).
Using microsensors and advanced bioimaging, we present
detailed data on the physical and chemical boundary conditions in
hospite and show how the metabolic activity of Prochloron and its
host modulate the microenvironmental conditions in response to
changes in irradiance. We discuss these new insights into the ecological niche of Prochloron and possible interactions with its host
and other microbes in light of its recently published genome and
recent studies of the overall microbial diversity and metagenome
of the didemnid ascidian host L. patella. (Donia et al., 2011a,b;
Behrendt et al., 2012a).
RESULTS
DISTRIBUTION AND PHOTOSYNTHETIC ACTIVITY OF PROCHLORON
Both macroscopic imaging and electron microscopy of L. patella
sections showed a dense colonization of the cloacal cavities and
the peribranchial space of zooids by apparently intact and dividing Prochloron cells (Figures 1 and 2A). Prochloron was closely
associated to the host tunic, either directly embedded in the tunic
or anchored in an exopolymeric substance apparent in the electron
micrographs (Figures 1D,E).
High absorptivity of red light was observed in the densely
pigmented Prochloron layer and the cyanobacterial biofilm underneath the ascidian, while more faint light absorption was observed
associated with the tunic matrix especially at the colony surface and immediately below the Prochloron layer (Figure 2B).
Prochloron exhibited a high maximum PSII quantum yield of ∼0.8
after dark acclimation in hospite, while the effective PSII quantum
yield declined down to ∼0.2 with increasing levels of incident
irradiance (Figure 2C). The cyanobacterial biofilm exhibited a
lower maximum PSII quantum yield of ∼0.5–0.6 decreasing to
an effective PSII quantum yield of <0.1 at the highest irradiance.
Using the product of absorptivity, quantum yield and incident
irradiance as a proxy for photosynthetic activity, the Prochloron
and cyanobacterial biofilm also showed different acclimation to
irradiance (Figure 2D), while more faint activity was found in a
thin layer on the tunic surface and a layer immediately below the
Prochloron cells. The cyanobacterial biofilm exhibited onset of saturation, as determined from the intersection of the initial slope and
the maximal activity level, at an irradiance of ∼100 µmol photons
m−2 s−1 , while the Prochloron layer showed saturation of photosynthesis at irradiances >250 µmol photons m−2 s−1 . Both zones
exhibited no photoinhibition due to short exposure to irradiances
of ∼1000 µmol photons m−2 s−1 . Imaging of the effective quantum yield of PSII and the derived relative photosynthesis revealed
further details on the distribution of photosynthetic activity in
L. patella (Figures 2E–L). The activity distribution in the cloacal
and peribranchial parts of L. patella closely followed the distribution of the green colored areas indicative of Prochloron and the
pink-colored cyanobacterial biofilm (compare, e.g., Figures 2A,I).
LIGHT PENETRATION AND ATTENUATION IN L. PATELLA
Scalar irradiance measurements in intact L. patella specimen
revealed both a pronounced scattering of incident light in the
upper tunic as well as strong attenuation of visible light (VIS;
Frontiers in Microbiology | Aquatic Microbiology
Prochloron microenvironment
400–700 nm) in the deeper layers (Figure 3A). VIS was strongly
scattered in the uppermost tunic leading to a local increase of scalar
irradiance reaching ∼140–170% of incident light. About 10–40%
of the incident VIS penetrated the upper tunic, while strong attenuation in the Prochloron-containing layer (ranging in depth from
∼1.5–2 to ∼4–6 mm below the tunic surface) reduced VIS down
to only a few % of the incident irradiance (Figure 3A). In the lower
part of the tunic, attenuation of VIS was less strong.
Near-infrared radiation (NIR) showed much less attenuation
but strong scattering in L. patella (Figure 3A). The scalar irradiance of NIR (700–750 nm) reached ∼170–230% of downwelling
NIR in the uppermost tunic. About 30–40% of the incident NIR
still prevailed below the Prochloron layer and NIR was less strongly
attenuated in the lower part of the tunic.
The scalar irradiance spectra showed further details on the
propagation and attenuation of light in L. patella (Figure 3B). In
the NIR region, weak minima around 800–805 and 870–880 nm
indicated the presence of some BChl a, while a stronger minimum
at 965–985 nm may indicate the presence of another BChl-like
photopigment. However, NIR was much less attenuated than visible wavelengths and combined with strong scattering this led
to significant local enhancement of the scalar irradiance in the
uppermost tunic.
In the visible spectral region, scalar irradiance showed strong
spectral minima corresponding to absorption maxima of several
photopigments: a strong minimum at 675 nm and a shoulder at
∼650 nm revealed the presence of Chl a and Chl b indicative of
Prochloron: a minimum at ∼625 nm, as well as minima at 585–
590, ∼565, and a shoulder at ∼490–495 nm indicated presence of
phycobiliproteins in L. patella. In the upper tunic, distinct minima
at ∼390 and ∼426 nm could indicate the presence of photoprotective pigments, but we did not confirm the type or quantity of
such compounds in this study.
DISTRIBUTION AND DYNAMICS OF O2
Profiling with thin O2 microelectrodes revealed a highly dynamic
microenvironment of Prochloron inside L. patella, which was
strongly regulated by ambient irradiance levels (Figure 4). In
the dark, only the uppermost part of the test remained oxic,
while the interior of L. patella was anoxic down to about 4 mm
depth into the ascidian colony, although the surface was exposed
to flowing aerated seawater in the flow chamber. Upon exposure to light, the ascidian showed rapid accumulation of O2 in
its interior cavities due to intense photosynthesis in the densely
populated Prochloron biofilm lining the peribranchial space and
the internal cloacal cavities, while the upper tunic exhibited O2
consumption both in darkness and light (Figure 4A). Under an
irradiance of 93 µmol photons m−2 s−1 , O2 started to accumulate
∼2 mm below the tunic surface, where the microsensor entered
the Prochloron layer; further, into this layer the declining O2 concentration indicated light limitation and net O2 consumption. At
higher irradiance O2 increased rapidly and approached saturating
levels reaching ∼250% air saturation in the Prochloron layer at
irradiances >300 µmol photon m−2 s−1 .
The O2 dynamics inside the ascidian was strongly affected by
irradiance. Experimental light-dark shifts with the O2 microsensors fixed ∼3 mm below the ascidian surface in the Prochloron layer
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A
Prochloron microenvironment
LISSOCLINUM
E
P A TE LLA
I
Quantum yield
Photosynthesis
ROCHLORON
CYA NOB A CTE RIA
B
Absorptivity
A=1-
C
ΦP =
1
F’M -F
F’M
PS = A·EPAR·ΦP
20 µmol photons m-2 s-1
J
F
RRED
RNIR
135 µmol photons m-2 s-1
PSII quantum yield
K
G
0.8
0.6
ROCHLORON
0.4
0.2
0
CYA NOB A CTE RIA
0
200
400
600
800
1000
290 µmol photons m-2 s-1
D
Relative photosynthesis
100
L
H
80
ROCHLORON
60
40
20
CYA NOB A CTE RIA
0
0
200
400
600
800
Irradiance (µmol photons m-2 s-1)
1000
460 µmol photons m-2 s-1
00
FIGURE 2 | Imaging of light absorption and relative photosynthetic
activity of Prochloron in hospite. (A) A cross-section of L. patella
showing a bright green layer of Prochloron cells in the cloacal cavities and
the peribranchial space of the zooids and a reddish cyanobacterial biofilm
colonizing the underside of the basal tunic. Light scattering spicules are
embedded in the tunic, especially in the upper tunic and immediately
below the Prochloron layer. (B) Absorptivity of red light (λmax 650 nm) in the
different layers of L. patella. (C,D) Quantum yield of PSII and derived
showed alternation from anoxic to hyperoxic conditions over time
scales ranging from 15 to 30 min, where the rate of O2 build-up
increased with irradiance indicative of an increased photosynthetic
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1
relative photosynthesis vs. irradiance curves for regions of interest (ROI) in
the Prochloron layer and the cyanobacterial biofilm colonizing the
underside of the ascidian. (E–H) Spatial distribution of PS II quantum yield
and the derived relative photosynthesis (I–L) in L. patella at selected
incident irradiance levels, E PAR . A proxy for the relative photosynthetic
activity was calculated from the relative PSII electron transport rate
rETR = ΦPSII E PAR , taking the different absorptivity over the imaged
specimen into account, as PS = A ΦPSII E PAR .
rate (Figure 4B). Even after prolonged time in darkness and
anoxia, Prochloron photosynthesis exhibited an immediate onset
of O2 production upon illumination. However, during profiling
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Kühl et al.
Prochloron microenvironment
A
A
B
B
FIGURE 4 | Oxygen dynamics in L. patella. (A) Depth profiles of O2
concentration (in % air saturation) at increasing irradiance levels (given as
numbers on curves in units of µmol photons m−2 s−1 ). (B) Dynamics of O2
concentration at ∼3 mm depth in a L. patella specimen under experimental
light-dark shifts at different irradiance levels (given as numbers on curves in
units of µmol photons m−2 s−1 ). Shaded areas indicate periods of
illumination.
FIGURE 3 | Light propagation in L. patella. (A) Depth profiles of
integrated scalar irradiance (in % of the incident downwelling irradiance at
the tunic surface) for VIS (VIS, 400–700 nm) and the part of near-infrared
radiation (NIR, 700–750 nm) which can potentially drive oxygenic
photosynthesis via Chl d or f. (B) Spectral scalar irradiance (in % of the
downwelling spectral irradiance at the tunic surface) measured at
increasing depth below the tunic surface (given as numbers on curves).
we often observed that the O2 level in the uppermost tunic of
L. patella decreased over time upon repeated profiling, especially
under high irradiance (data not shown), and such apparent wound
reactions caused long term downward drift in many measurements
and limited the amount of O2 profiles obtained.
While the O2 microsensors only provided a limited number of
spot measurements in the ascidians (Figure 4), the O2 dynamics
were confirmed by imaging of O2 concentration across a L. patella
specimen sliced vertically and pressed up against an O2 sensitive
foil mounted inside the transparent flow chamber (see Materials
and Methods for further details). This O2 imaging revealed similar dynamics in L. patella as observed in the microsensor data,
showing anoxia in the Prochloron layer in darkness and enhanced
O2 due to onset of photosynthesis upon illumination (Figure 5).
However, the actinic light source used in the imaging setup only
allowed homogenous illumination of the ascidian with a maximal
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photon irradiance of 250 µmol photons m−2 s−1 and this did not
enable high super saturating O2 levels to be reached.
Oxygen imaging revealed several zones in the ascidian contributing to the build-up of O2 (Figures 5C–E). Initially and
shortly after onset of illumination, O2 evolution was detected in
separate zones comprising the upper tunic surface, the Prochloron
layer and the cyanobacterial biofilm layer on the ascidian underside
(Figure 5D). As the O2 levels continued to increase and oxygenate
the ascidian tissue, these distinct zonations between the test surface
and the Prochloron layer became less distinct, while a second zone
of O2 production just below the Prochloron layer and above the
cyanobacterial biofilm became visible (Figures 5F,G). Upon darkening, the Prochloron layer exhibited the fastest O2 depletion, while
the other regions in L. patella approached anoxia more gradually
(Figures 5I–L). The upper tunic and the biofilm on the underside of L. patella remained oxic in darkness, albeit at low levels of
∼10–20% air saturation (Figure 5; ROI 1 and 6 in Figure 6).
DISTRIBUTION AND DYNAMICS OF PH
pH is another chemical parameter in the ascidian host that potentially is strongly affected by Prochloron photosynthesis due to their
CO2 -fixation. Attempts to profile pH with glass needle microelectrodes were not successful due to their fragility and apparent local
release of acid vacuoles in the ascidian tunic when inserting the
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Prochloron microenvironment
FIGURE 6 | Dynamics of O2 concentration in selected regions of
interest (ROI; see Figure 5L) after on set of illumination with a photon
irradiance of 250 µmol photons m−2 s−1 and after darkening. Symbols
represent the average ± standard deviation in each ROI.
FIGURE 5 | Imaging of O2 concentration dynamics in L. patella. (A–J)
Pseudocolor images of the O2 distribution in L. patella under an incident
photon irradiance of 250 µmol photons m−2 s−1 and after darkening. Times in
light and after darkening are indicated in minutes in each panel. (K)
Photograph of L. patella cross-section. (L) Image of cross-section
illuminated with red light showing the Prochloron layer and the biofilm
underneath the ascidian. Regions of interest (ROI) show the origin of data
plotted in Figure 6.
Layers in the upper tunic (e.g., ROI2 in Figure 7, monochrome
image) and the basal tunic (e.g., ROI3 and ROI5 in Figure 7, monochrome image) showed slightly acidic conditions around pH 6.5–7
both in darkness and during illumination. A patch of cyanobacterial biofilm colonizing the underside of L. patella (ROI4 in Figure 7,
monochrome image) showed a slightly higher pH reaching pH
7.5–7.8 (with the exception of one time point showing an intermittent dip to pH 7.0), and exhibited no clear light-dark dynamics
under the experimental irradiance levels.
The pH images showed some conspicuous hot spots of strong
pH dynamics in part reflecting different amounts of Prochloron
in the cloacal cavities (Figure 7). However, the surrounding host
tissue also exhibited an interesting pattern of fluctuating pH, e.g.,
initially becoming more acidic upon onset of illumination and
then exhibiting fluctuating yet acidic pH conditions (Figure 7,
light on 0–60 min). Such fluctuations and hot spots were also evident after darkening, where the host tissue in some regions of
the upper test apparently became less acidic (Figure 7, light off
10–30 min).
relatively large needle electrodes (data not shown). Therefore, we
adapted a new pH imaging approach to resolve the pH dynamics
in L. patella. As this is the first time such imaging has been applied,
detailed data on sensor performance, and calibration are given in
the methods section at the end of this article.
The pH dynamics in L. patella showed pronounced spatiotemporal pH variations and distinct differences to the pH in the
surrounding seawater (Figures 7 and 8). The layers containing
Prochloron in the peribranchial space and common cloacal cavity
(e.g., ROI 1 in Figure 7, monochrome image) showed the most
pronounced pH dynamics with irradiance, shifting from about
pH 7 to 7.5 in darkness to almost pH 10 under an irradiance of
250 µmol photons m−2 s−1 . This shift occurred within ∼20 min
in response to a light-dark shift.
Application of microsensors and bioimaging revealed new and,
up to now, the most detailed insights to the microenvironment
of Prochloron and its host L. patella and confirmed preliminary
data obtained in the didemnid ascidian Diplosoma virens (Kühl
and Larkum, 2002) and the first few light and O2 measurements
in L. patella obtained during a survey of its microbiome (Behrendt
et al., 2012a). In addition to more detailed microsensor measurements of O2 and scalar irradiance, this study presents the
first use of O2 and pH imaging in photosymbiotic ascidians and
combines this new methodology with variable chlorophyll fluorescence imaging of light absorption and PSII activity at similar
high spatio-temporal resolution. This allowed us to map both
the dynamic chemical landscape and the distribution of photosynthetic activity onto the structural heterogeneity of L. patella
DISCUSSION
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November 2012 | Volume 3 | Article 402 | 7
Kühl et al.
FIGURE 7 | Pseudocolor images of the pH distribution in L.
patella under an incident photon irradiance of
250 µmol photons m−2 s−1 and after darkening. Times in light and
after darkening are indicated in minutes in each panel. The
Prochloron microenvironment
monochrome image (lowermost right panel) is the cross-section
illuminated with red light showing the Prochloron layer and the
biofilm underneath the ascidian. Regions of interest (ROI) are outline
din blue and show the origin of data plotted in Figure 8.
and its photosymbionts. In the following, we discuss the implications of these new microenvironmental analyses in light of what
is currently known about the physiology and diversity of photosymbionts in the Prochloron-didemnid ascidian association and its
associated microbiome.
OPTICAL PROPERTIES AND LIGHT PROPAGATION
Microscale light measurement showed that both visible and NIR
were scattered strongly in the opaque tunic of L. patella and such
local photon trapping led to local maxima in scalar irradiance
significantly above the incident downwelling irradiance, i.e., up
to 170 and 230% for VIS and NIR, respectively. Similar photon
trapping has been observed in different optically dense biological systems with pronounced scattering like sediments, biofilms,
plant, and animal tissue (e.g., Vogelmann and Björn, 1986; Vogelmann, 1993; Kühl et al., 1994, 1995; Magnusson et al., 2007). The
magnitude of the scalar irradiance maximum is modulated by the
optical properties of the system, especially the refractive index, the
absorption and scattering coefficients as well as the characteristic
scattering phase function of the matrix (see details in Kühl and
Jørgensen, 1994).
The optical properties of the tunic matrix are largely unknown
but with the exception of the genus Diplosoma, photosymbiotic
didemnids are known to contain scattering spheroid spicules in
their tunic. It has been speculated that these calcareous spicules
Frontiers in Microbiology | Aquatic Microbiology
FIGURE 8 | Dynamics of pH in selected regions of interest (ROI; see
Figure 7) after on set of illumination with a photon irradiance of
250 µmol photons m−2 s−1 and after darkening. Symbols represent the
average ± standard deviation in each ROI.
could have an important role for light propagation and photoadaptation in the tunic of photosymbiotic ascidians (Hirose et al.,
2006). A role for spicules in light guiding has been demonstrated
November 2012 | Volume 3 | Article 402 | 8
Kühl et al.
in sponges harboring photosymbionts, where radially arranged
bundles of spicules channel light into deeper layers of the sponge
(Brümmer et al., 2008). We did not see any evidence for light
guiding in L. patella when imaging cross-sections of a colony, while
irradiating it from above. However, it is known from other tissues,
such as leaves (Gorton et al., 2010) and corals (Wangpraseurt et al.,
2012), that direct and diffuse light can have very different results
on light scattering in internal tissues.
Moreover, spicules in photosymbiotic ascidians do not form
bundles and can be regarded amorphous scattering particles distributed in the tunic – often at higher density in the upper layers.
Hirose et al. (2006) studied the Prochloron-containing ascidian
Didemnum molle, which showed a decreasing density of spicules
in the tunic with increasing water depth, and they proposed that
high spicule density in shallow water primarily induced photoprotection by increasing the reflectivity of the ascidians. We are
not aware of similar studies of the spicule density in L. patella,
but our light measurements showed pronounced scattering in the
upper tunic leading to photon trapping and local amplification of
scalar irradiance, especially in the NIR region, where absorption
in the tunic is minimal and such scattering effects therefore are
most pronounced (Figure 3).
Increased photon pathlength due to spicule-induced scattering
in the tunic is but one of several mechanisms potentially affecting
light levels and photon trapping. The refractive index of the tunic
matrix relative to that of the surrounding seawater can also play
an important role as refractive index mismatches can enhance
redirection of upwelling scattered light back into the tunic and
further enhance photon pathlengths of scattered light (Kühl and
Jørgensen, 1994).
A somewhat similar mechanism, assuming absence of scattering in the tissue layer, has been proposed in corals, where multiple
scattering and enhanced diffuse reflectance from the coral skeleton underneath leads to a more efficient absorption of incident
light by photopigments (Enriquez et al., 2005). In the case of completely diffuse backscatter, the average path length of upwelling
photons is twice that of photons in a collimated light beam traversing a thin layer of tissue (Kühl and Jørgensen, 1994). This
implicates that the probability for a photon to become absorbed is
significantly enhanced, as long as the density of absorbers is not so
high that self-shadowing and package effects arise. As a result of
this phenomenon, corals inhabiting high light environments can
maximize their absorption capacity with low pigment investment
while reducing self-shading (Enriquez et al., 2005). Similar effects
may be at play in the spicule-containing tunic of photosymbiotic
ascidians.
All the above mentioned light trapping and scattering mechanisms lead to enhanced spectral filtering as an increased photon
path length due to scattering increases the probability for encountering absorbing pigments in the tunic. Accordingly, photons at
wavelengths within major absorption bands of host and photosymbiont pigments will be absorbed more efficiently, while
photons outside such absorption bands will be unaffected leading to larger relative differences in the light spectrum. This can
have important implications for the Prochloron-ascidian association, where UV-screening mycosporine like aminoacids (MAAs)
have been found in the upper tunic (e.g., Hirose et al., 2004).
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Prochloron microenvironment
Genomic analyses show that such compounds can be synthesized
by Prochloron (Donia et al., 2011b) but the translocation to the
ascidian host tissue remains to be studied, and Prochloron may not
be the only source. We hypothesize that the presence of spicules
and the scattering properties of the upper tunic enable efficient
UV protection in analogy to similar effects in corals (Enriquez
et al., 2005), while at the same time ensuring that sufficient visible wavelengths can propagate into deeper layers with Prochloron.
Interestingly, we also found a higher density of spicules in the
tunic immediately below the Prochloron layer, which may further
affect the light field and spectral absorption in L. patella, but more
detailed optical studies are needed to test such hypotheses.
IN VIVO CONDITIONS AND PHOTOSYNTHESIS OF PROCHLORON
Our microenvironmental measurements show that Prochloron,
in the peribranchial space and cloacal cavity, inhabits a very
dynamic ecological niche in L. patella, wherein chemical conditions respond rapidly to changes in irradiance over a timescale
of 15–30 min. In the case of O2 , levels changed quickly from
anoxic in the dark up to >250% O2 air saturation at PAR irradiances >300 µmol photon m−2 s−1 (Figure 4B). The O2 imaging
approach supported these results and in addition allowed a much
more detailed assessment of the contributions of other oxygenic
photosymbionts in L. patella.
The pH imaging system revealed that the pH in the peribranchial/common cloacal cavity shifted from ∼pH 7.0 to pH
7.5 in the dark to just below pH 10.0 under a PAR irradiance of
∼250 µmol photon m−2 s−1 . The pH in this zone is thus lower
than in seawater (∼pH 8.2) in the dark and much higher under
moderately high irradiance. As the peribranchial/common cloacal cavities are connected to the external seawater by inhalant
siphons, one per branchial cavity, it is clear that the internal
environment is strongly influenced by the ascidian tissues and
Prochloron both in the dark and in the light. While photosynthesis can proceed at >pH 9.5 (e.g., Giordano et al., 2005)
it is entirely dependent on HCO−
3 at that pH, since CO2 is
absent under such alkaline conditions. This suggests that inorganic carbon transport in Prochloron is mainly HCO−
3 -dependent.
This conclusion is consistent with the evidence from a recent
genomics study (Donia et al., 2011b) showing that Prochloron
only has low-affinity carbon transport pumps. Prochloron also
lacks many transporters found in other cyanobacteria, including
those involved in high-affinity CO2 uptake (NdhD3-F3) and the
low-CO2 inducible bicarbonate transporters sbtA and CmpA-D
(Badger et al., 2002). Interestingly, the upper and lower tunic layers that showed some evidence of O2 -evolving cyanobacteria were
acidic in the light, possibly reflecting a much greater influence of
surrounding ascidian cells. The slightly acidic tissue adjacent to the
peribranchial/common cloacal cavities and the local hotspots also
reflected changes in the animal host tissues, probably in response
to the massive change in pH of the peribranchial/common cloacal
cavities.
The data presented here for the first time show in vivo
photosynthesis of Prochloron in the peribranchial/cloacal cavity to be both high and efficient (Figures 2–4). Early work
on freshly extracted Prochloron (Critchley and Andrews, 1984)
indicated that photosynthetic O2 evolution was very efficient
November 2012 | Volume 3 | Article 402 | 9
Kühl et al.
[up to 180 µmol O2 (mg Chl)−1 h−1 ] under irradiances of 300–
400 µmol photons m−2 s−1 , supported by in vivo measurements
of variable chlorophyll fluorescence (Schreiber et al., 1997). The
direct measurements here are similar (Figure 2) and the light
measurements suggest that in shallow reef waters Prochloron can
experience irradiance levels of >300–400 µmol photons m−2 s−1
in hospite. It is noteworthy that in our variable chlorophyll fluorescence imaging, we used blue actinic light exposing a cut surface
of L. patella evenly, so that the filtering effect of the upper tunic
was avoided. The apparent lack of photoinhibition indicates that
even at higher irradiance photosynthesis would not be impaired.
Our data are in line with earlier observations on high irradiance
tolerance of photosynthesis in Prochloron when associated with its
didemnid host (e.g., Alberte et al., 1987; Lewin and Cheng, 1989).
The presence of UVA/B absorbing MAAs in the outer tunic, shown
in previous work (e.g., Hirose et al., 2004) and by the in vivo spectra
here (Figure 3B), could be an important factor allowing high photosynthetic rates of Prochloron in situ even at high ambient solar
irradiance. Furthermore, Lesser and Stochaij (1990) demonstrated
the presence of antioxidant enzymes such as superoxide dismutase, ascorbate peroxidase, and catalase in the Prochloron-L. patella
symbiosis, and showed that the activity of these enzymes were
directly proportional with irradiance. Prochloron thus seems well
adapted to quench ROS formed in the photosynthetic apparatus
under high irradiance. This is further supported by the presence
of multiple hli genes in the genome of Prochloron (Donia et al.,
2011b). These genes encode so-called high light induced proteins
that are thought to play a role in quenching of excess excitation
energy (He et al., 2001).
As pointed out in the above section, under these high photosynthetic conditions, the O2 concentrations and pH in the
peribranchial/cloacal cavity rise to very high levels (Figures 5–9).
Clearly neither of these factors inhibits the photosynthetic capacity
of Prochloron and as pointed out above, the low-affinity inorganic carbon transporters in the plasma membrane of Prochloron
must be able to transport adequate quantities of HCO−
3 into the
cells. Whether any role is played by matrix compounds present in
the Prochloron layer (Figure 1), and whether the host exerts any
control on the uptake of inorganic carbon across the plasma membrane (Critchley and Andrews, 1984; Griffiths, 2006) are important
questions for future research.
OTHER OXYPHOTOTROPHS IN DIDEMNID ASCIDIANS
Besides Prochloron, other phototrophs occur in the upper unic of L.
patella and as judged from the in vivo spectral light measurements
(Figure 3) appear to be phycobilin-containing cyanobacteria. This
is in line with the findings in a recent survey of the microbial diversity in L. patella, where 16S RNA gene sequences and
hyperspectral signatures of cyanobacteria were abundant in the
upper tunic (Behrendt et al., 2012a). Another cyanobacterial zone
also lies in the tunic beneath the Prochloron layer, the photosynthetic activity of which became evident in the imaging of PSII
activity, O2 and pH. The slower dynamics of O2 in this layer
is probably the result of light screening by cyanobacteria in the
upper test and the Prochloron layer. Both zones of cyanobacteria
have been observed before and the tunic is known to harbor the
cyanobacterium Synechocystis trididemni and other cyanobacteria
Frontiers in Microbiology | Aquatic Microbiology
Prochloron microenvironment
A
B
FIGURE 9 | Spectral characteristics and calibration of sensor foils used
for pH imaging. (A) Spectral properties of the pH sensor material showing
emission spectra of the pH planar optode (λexc = 450 nm, 0.6 M NaCl, 25˚C)
at varying pH (indicated on curves), chemical structures of the dyes and the
windows (W1 and W2) used for the interrogation with a multispectral
imaging system. The spectra were obtained on a Hitachi F-7000
fluorescence spectrometer. (B) Calibration curves of the pH sensor foil, i.e.,
a plot of the ratio of W2/W1 vs. defined pH levels. Note that the optimal
dynamic range of the optode is at pH 6–9. The sigmoidal calibration curve
flattens at pH >9 and <6, and pH values obtained in these ranges are thus
less accurate.
with special pigmentation (Lafargue and Duclaux, 1979; Kott,
1984; Cox et al., 1985; Hirose et al., 2009b; López-Legentil et al.,
2011). López-Legentil et al. (2011) found Chl d-containing Acaryochloris cells in the unic of other didemnid ascidians on mangrove
roots in the Bahamas but we have not found any evidence for Chl
d in the unic of L. patella (Kühl et al., 2005; Behrendt et al., 2012a).
The other oxyphototrophs in L. patella are exposed to very
different irradiance regimes in comparison to Prochloron. Those
colonizing the surface and upper tunic would be exposed, at times,
to high and potentially photoinhibitory irradiance levels reaching >400 µmol photons m−2 s−1 . In contrast, S. trididemni and
other cyanobacteria below Prochloron layer would be much better
shielded and prone to light limitation at high irradiance as a result
of the strongly absorbing Prochloron layer above (Figure 3). Due to
strong light attenuation and spectral filtration by Prochloron, the
cyanobacteria thriving below it have adapted their pigmentation
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Kühl et al.
and S. trididemni and other cyanobacteria present contain phycourobilin in addition to other more conventional phycobiliproteins (Cox et al., 1985; Hirose et al., 2009b), which would allow
them to harvest green light very efficiently; it is probably phycourobilin that contributes to the high absorption at around 500 nm in
the deeper tunic layers (Figure 3B; cf Hirose et al., 2009b). In
the present study, we did not specifically target the identification
and abundance of these cyanobacteria in the tunic of L. patella.
Behrendt et al. (2012a) also did not extend their microbial diversity
survey to the inhabitants of the lower tunic in L. patella; however,
our data point to a rather dense and active community, which
awaits further investigation.
Our imaging studies also revealed the presence of oxyphototrophs in the biofilm layer colonizing the underside of L. patella.
This biofilm is highly diverse, with cyanobacteria as the predominant oxyphototrophs, especially A. marina that thrives in this
ecological niche deprived of visible wavelengths but with abundant NIR supporting its Chl d-based oxygenic photosynthesis
(Kühl et al., 2005; Behrendt et al., 2012a; Larkum et al., 2012).
The present results thus confirm our previous studies (Kühl et al.,
2005, 2007a) showing active photosynthesis in this biofilm albeit
with a more moderate dynamics in both O2 and pH as compared
to Prochloron. Actually, both the O2 and pH imaging data represent the very first in vivo measurements of these parameters in
the natural habitat of A. marina, but a detailed discussion of the
implications of these observations for our knowledge on the in situ
biology of Chl d-containing cyanobacteria is beyond the scope of
this study.
THE MICROBIOME OF L. PATELLA
Sequencing of the 16S rRNA gene and bulk metagenomic analysis have provided detailed information on the composition of
a tremendous variety of microbial assemblages, ranging from
the human body to marine environments. Similar studies performed on ascidian-associated microbial communities revealed a
high bacterial diversity (Martínez-García et al., 2007; Donia et al.,
2011a; Behrendt et al., 2012a), possibly due to distinct microenvironments as demonstrated in this study, and domination by
selected phyla such as proteobacteria and cyanobacteria (Tait et al.,
2007; Menezes et al., 2010; López-Legentil et al., 2011; Behrendt
et al., 2012a). Combined molecular and microenvironmental measurements on L. patella demonstrated the coexistence of three very
different microbial communities, separated by only a few millimeters of animal tissue (Behrendt et al., 2011): (i) a biofilm on the
upper surface exposed to high irradiance, (ii) a cloacal cavity dominated by Prochloron spp. characterized by strong depletion of VIS
and a dynamic chemical microenvironment, and (iii) a biofilm
community covering the underside of the animal, where light is
depleted of visible wavelengths and enriched in NIR. Further molecular analysis of the L. patella microbiome revealed bacteria with
a rich genetic diversity, capable of producing significant amounts
of secondary bioactive metabolites (Donia et al., 2011a) and earlier
studies highlighted Prochloron as one of the key producers of such
compounds (Schmidt et al., 2005). Further metagenomic investigations of Prochloron cells revealed an arsenal of functional gene
cassettes coding for secondary metabolites across large geographical distances, with surprisingly little genomic diversion within
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Prochloron microenvironment
the different Prochloron cells (Donia et al., 2011a). Even though
Prochloron seems to functionally depend on its host, its genome
contains a set of core genes similar to other cyanobacteria (Donia
et al., 2011b), which, in all likelihood allows life outside of the
ascidian-Prochloron association. Despite this, larger amounts of
free-living Prochloron cells have only been reported once (Cox,
1986), while facultative relationships have been reported between
non-ascidian invertebrates and Prochloron cells (Cheng and Lewin,
1984; Parry, 1986).
The abundance of Prochloron cells appears to underlie seasonal
changes in temperature (McCourt et al., 1984) and more generally
the distribution of cyanobacteria is found to be governed by the
composition and availability of light (Stomp et al., 2007a,b). NIRutilizing bacteria associated with ascidians are a prime example of
such a highly specialized niche-partitioning: the Chl d-containing
cyanobacterium A. marina was found to survive below didemnid
ascidians in a light environment relatively enriched in NIR and
depleted in VIS due to the overlying layer of Chl a/b-containing
Prochloron cells (Kühl et al., 2005, 2007a; Behrendt et al., 2012a).
Such microenvironmental patterns can even be extrapolated to
larger biological frameworks: the sampling of L. patella associated
biofilms along a depth gradient thus revealed a negative correlation between the abundance of NIR-utilizing phototrophs (A.
marina, Rhodospirillaceae, Rhodobacteraceae, Chloracidobacteria)
and water depth indicating that their abundance was strongly
influenced by the availability of NIR, which is readily attenuated by
seawater (Behrendt et al., 2012a). This highlights the need to perform ecologically relevant molecular surveys of microbial diversity
on an appropriate (micro-) scale with accompanying metadata.
In this context, microenvironmental data can inform both observations obtained by genomics and even the actual planning of
such surveys. We have demonstrated that Prochloron lives in a
niche experiencing strong fluctuations of the chemical microenvironment, generally experiencing hypoxia or anoxia inside their
ascidian host under low light or dark conditions. Such microenvironmental settings thus point to a potential presence of a number
of anaerobic processes and microbes.
In densely populated cyanobacterial communities experiencing periods of hypoxia/anoxia, N2 fixation is often found (Stal and
Zehr, 2008; Steunou et al., 2008). Diazotrophy has been reported
several times in photosymbiotic ascidians, but whether Prochloron
can fix N2 remains unclear (Paerl, 1984; Koike et al., 1993; Kline
and Lewin, 1999). Interestingly, genes associated with N2 fixation were absent in the recently published Prochloron genome,
apparently forcing it to rely on recycled nitrogen from its host
via a number of nitrogen transformation pathways present in the
genome (Donia et al., 2011b). This fits with the observation that
ammonium is the major nitrogenous waste product of the ascidian host (Goodbody, 1974), which is also effectively taken up by
Prochloron (Parry, 1985), while the presence of genes for urease
and urea uptake point toward a potential role of urea in the nitrogen cycling between host and symbiont (Donia et al., 2011b).
Diazotrophy may, however, occur in Prochlorons microenvironment as other N2 -fixing bacteria such as Azospirillum brasilense
were found to reside in the cloacal cavity of L. patella (Behrendt
et al., 2012a) and this may explain some of the earlier observations of N2 fixation (Paerl, 1984). Nevertheless, Prochloron is
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Kühl et al.
clearly the predominant inhabitant of the peribranchial space and
inner cloacal cavities of L. patella, and we speculate that such diazotrophs may benefit from the strong O2 dynamics imposed by
Prochloron.
It remains to be investigated what type of dark metabolism
Prochloron is thriving on during periods of low or no light and
anoxia. Our data show that oxic dark respiration of Prochloron
is limited due to strong O2 depletion in its microenvironment.
Under such conditions, cyanobacteria can rely on fermentation
for their energy metabolism (Stal and Moezelaar, 1997). To our
knowledge, the presence and activity of fermentative pathways
in Prochloron have not been investigated. A first look into the
Prochloron genome published by Donia et al. (2011b) actually
revealed the potential presence of both mixed acid as well as lactate/pyruvate fermentation pathways (data not shown), which are
known to occur in cyanobacteria (Stal and Moezelaar, 1997). Our
finding of low pH in darkness support the speculation that fermentation may be an important part of Prochloron dark metabolism.
Confirmation of this and further insight to the dark metabolism
of Prochloron awaits more dedicated physiological studies of its
carbon metabolism, preferentially in combination with further
scrutiny of the Prochloron genome and subsequent transcriptome
analyses. The same holds true for other aspects of Prochloron
metabolism and biosynthesis, such as the synthesis of UV protective MAAs, abundant lipids, sterols, and bioactive secondary
metabolites in L. patella (Donia et al., 2011a,b). While the presence of these pathways in Prochloron has been shown, we still
know nothing about the diel dynamics of gene expression and
enzyme synthesis in photosymbiotic ascidians and how particular
microenvironmental controls play a regulatory role. A combination of microenvironmental analysis with genomics approaches
(see, e.g., Steunou et al., 2008; Jensen et al., 2011) to further resolve
the biology of Prochloron thus seems a very promising way to
proceed.
Prochloron remains uncultivated despite numerous attempts
since its discovery in 1975 (Hirose et al., 2009a). An hitherto unconfirmed report of successful cultivation argued that
Prochloron lacked the ability to synthesize tryptophan (Patterson and Withers, 1982), while the recent Prochloron genome
shows the presence of a full set of tryptophan anabolic genes
(Donia et al., 2011b). Interestingly, Patterson and Withers (1982)
also found cell division of Prochloron under acidic conditions with maximal growth at pH 5.5 under low irradiance of
∼110 µmol photons m−2 s−1 over a 18:6 h light-dark cycle. We
found acidification of the Prochloron microhabitat in response
to darkness, albeit only a few areas reached pH <6 (see, e.g.,
Figures 7C,D). In contrast, Prochloron photosynthesis is limited
to a higher pH range with a maximum at pH 8–8.5 (DionisioSese et al., 2001). Not much is known about the cell division cycle of Prochloron, besides one study reporting a diurnal rhythm with maxima of dividing cells in early morning
and afternoon (Lewin et al., 1984), and we do not know how
such cell division pattern is modulated by microenvironmental
changes.
With the new knowledge on pH and O2 dynamics presented
here, it is possible to further optimize enrichment and cultivation scenarios better mimicking the natural habitat of Prochloron
Frontiers in Microbiology | Aquatic Microbiology
Prochloron microenvironment
and to test new working hypotheses such as the importance of
chemical and light gradients for Prochloron growth, and the importance of keeping Prochloron in a biofilm/aggregated growth mode
for longer survival outside its host. A promising approach for
enrichment and growth of Prochloron cells in gradients under controlled biofilm-like conditions could be based on immobilization
of extracted Prochloron cells in alginate beads; an approach recently
shown to work with A. marina (Behrendt et al., 2012b).
MATERIALS AND METHODS
FIELD SITE AND SAMPLE COLLECTIONS
Didemnid ascidians were sampled at low tide on the outer reef flat
and crest off Heron Island (S23˚26′ 0055, E151˚55′ 0850), Great
Barrier Reef in the Austral summer (January–February) during several field trips in the period 2001–2012. Specimen of L.
patella were sample as larger intact specimens (5–20 cm2 ; 5–
20 mm thick) covering coral patches on the outer reef flat and
down to ∼4 m depth on the reef crest (Figure 1). A more detailed
description of sampling and field sites is given in Behrendt et al.
(2012a).
Collection of didemnids was either done by hand or snorkeling. The samples were immediately transported in a bucket
with sea water back to Heron Island Research Station, where they
were transferred to outdoor aquaria continuously flushed with
fresh aerated seawater (26–28˚C) pumped in from the reef. Frequent exchange of water by flushing and continuous strong mixing
of water in the aquarium is necessary to avoid degradation of
the ascidians. To avoid high solar irradiance, the aquaria were
covered by shading cloth dampening solar irradiance to ∼200–
300 µmol photons m−2 s−1 . Under such conditions we could keep
the sampled ascidians healthy and with actively photosynthesizing
Prochloron for up to a week.
Measurements were done on small intact colonies of L. patella
(∼2–4 cm2 , 5–10 mm thick), as well as on specimens with an
exposed fresh vertical cut through the test or, alternatively, on
1–2 mm thick vertical sections of L. patella. Transfer, slicing and
cutting of specimen were done in a large beaker of seawater to
buffer pH changes due to release of acid vacuoles. Relatively flat
and homogeneous pieces of L. patella with a surface area of a few
cm2 were cut with a scalpel and immediately rinsed and submerged
in filtered seawater. Cross-sections were cut from homogenous
pieces with a razor blade for subsequent imaging. Generally, specimens showed a high maximal PSII quantum yield of >0.7 for
several hours after such handling indicative of fast recovery and
minimal stress on Prochloron.
ELECTRON MICROSCOPY
TEM
Specimens were fixed with 2% v/v glutaraldehyde in 0.05 M
sodium phosphate buffer (pH 7.2). Following isolation of suitable specimen blocks, the samples were rinsed three times in
0.15 M sodium cacodylate buffer (pH 7.2) and subsequently postfixed in 1% w/v OsO4 in 0.12 M sodium cacodylate buffer (pH
7.2) for 2 h. The specimens were dehydrated in graded series of
ethanol, transferred to propylene oxide, and embedded in Epon
according to standard procedures. Sections, ∼80 nm thick, were
cut with a Reichert-Jung Ultracut E microtome and collected on
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Kühl et al.
copper grids with Formvar supporting membranes. Ultra thin
sections were collected on copper grids with Formvar supporting
membranes and stained with uranyl acetate and lead citrate, and
subsequently examined with a Philips CM 100 TEM (Philips, Eindhoven, Netherlands), operated at an accelerating voltage of 80 kV
and equipped with an OSIS Veleta digital slow scan 2k × 2k CCD
camera. Digital images were recorded with the ITEM software
package.
FIB (focused ionbeam) SEM
Specimens were fixed with 2% glutaraldehyde in 0.05 M phosphate buffer (pH 7.2) and postfixed in 1% w/v OsO4 with 1.5%
potassium ferrocyanide. Following a rinse in dH2 O the specimens
were stained en bloc in 1% Uranyl acetate over night, dehydrated
in ethanol, and embedded in Epon according to standard protocols. The Epon blocs were mounted on aluminum stubs with
colloidal carbon as an adhesive, sputter-coated with gold (Polaron
SEM Coating Unit E5000), and imaged with a Quanta 3D FEG
(FEI) operated at 5 kV using a vCD backscattered electron detector. Additionally, mounted cross-sections of the same samples were
imaged on a Tecnai G2 20 Twin transmission electron microscope.
IMAGING OF VARIABLE CHLOROPHYLL FLUORESCENCE
Cross-sections through L. patella were placed in a small Petri-dish
and covered by a thin (∼1–2 mm) layer of seawater and investigated with a variable chlorophyll fluorescence imaging system
(I-PAM, Walz GmbH, Germany) consisting of a CCD camera, a
LED ring light (with blue, red, and NIR LEDs), and a controlling
unit connected to a PC running the dedicated imaging system software (Imaging-WIN 2.3, Walz GmbH, Germany). The blue LEDs
(470 nm) provided both weak pulses of measuring light as well
as defined levels of actinic light measured as downwelling photon
irradiance, E d , in the focus plane of the system with a calibrated
irradiance meter (LI-250 and LI-192, LiCor, USA). A detailed
description of variable chlorophyll fluorescence imaging systems
is given elsewhere (Grunwald and Kühl, 2004; Ralph et al., 2005;
Kühl and Polerecky, 2008; Trampe et al., 2011). The system enabled
quantification of a proxy for PAR absorptivity, A, by measuring the
reflected red light (R, 650 nm) and NIR (R NIR , 780 nm) from the
specimen/sample and calculating the ratio A = 1 − (R/R NIR ).
Based on the non-actinic imaging of the chlorophyll fluorescent
yield before and during a strong saturation pulse, using weak modulated blue measuring light, several parameters characterizing the
distribution and photosynthetic performance of oxyphototrophs
in the ascidian specimens could be quantified and visualized. This
included:
(1) The maximal quantum yield of PSII electron transport,
Φmax = (F m -F 0 )/F m, where F0 is the minimal fluorescence
yield and F m is the maximal fluorescent yield of the darkadapted sample measured prior to and during a strong
saturation pulse, respectively;
(2) The effective quantum yield of PSII electron transport
′ − F )/F ′ , where F is the fluorescent yield
ΦPSII = (Fm
m
′ is the maxunder a known level of blue actinic light and Fm
imal fluorescent yield during a subsequent saturation pulse,
respectively.
www.frontiersin.org
Prochloron microenvironment
Based on these measurements, we calculated images of a range
of derived photophysiological parameters characterizing photochemical and non-photochemical quenching of absorbed light
energy in Prochloron and other oxyphototrophs present in the
ascidians: a proxy for the relative photosynthetic activity was calculated from the relative PSII electron transport rate rETR = ΦPSII
E d by taking the different absorptivity over the imaged specimen
into account, as PS = A ΦPSII E d .
The imaging system enabled acquisition of the mentioned parameters under a range of different actinic light levels applied to the
samples over pre-defined time periods ranging from 10–20 s to
10 min. In this way, we could obtain information on both rapid and
slow responses of photosynthetic performance. So-called rapid
light curves (RLC) were obtained my measuring ΦPSII at increasing irradiance using 10–20 s exposure to each irradiance level. Such
light curves provide a snap-shot of the current photosynthetic
capacity for handling light but should not be regarded similar to
traditional photosynthesis vs. irradiance curves that are measured
at steady state after longer incubation times (Schreiber et al., 1997).
Such steady state light curves (SLC) of ΦPSII vs. irradiance were
obtained using 5–10 min long incubation times. Besides images,
information averaged over particular regions of interest (ROI) was
extracted with the system software (Image WIN 2.3, Walz GmbH,
Germany).
MICROSCALE LIGHT MEASUREMENTS
Light propagation in the ascidian tissues was measured with
fiber-optic scalar irradiance microprobes (Kühl, 2005) coupled
to a fiber-optic spectrometer (QE65000, Ocean Optics, Dunedin,
USA) for measurements of spectral scalar irradiance. Intact ascidian samples were fixed onto a black neoprene holder by thin
insect preparation needles and placed in a flow chamber. The
scalar irradiance microprobe was mounted in a manually operated micromanipulator (MM33, Märzhäuser, Wetzlar, Germany)
and inserted into the ascidian tissue at an angle of 45˚ relative
to the vertically incident light from a fiber-optic halogen lamp
equipped with a collimating lens (KL2500, Schott, Germany). The
solid test material of L. patella did not allow direct insertion of the
spherical microprobe tip, and it was necessary to make a minute
incision with the tip of a thin (29G) hypodermic needle prior to
measurements.
All scalar irradiance measurements were normalized to the
incident downwelling irradiance as measured with the scalar irradiance microprobe positioned in a black light well at a similar
position in the light field as over the unic surface. Spectral measurement in different depths were also integrated over visible
wavelengths (400–700 nm) driving oxygenic photosynthesis in
Prochloron as well as over a region in the NIR (700–740 nm) that
can be used by oxyphototrophs containing Chl d or f. This allowed
calculation of depth profiles of VIS and NIR scalar irradiance in %
of the incident downwelling irradiance in the respective spectral
ranges.
MICROSCALE O2 AND PH MEASUREMENTS
Microscale O2 measurements were done with amperometric
Clark-type O2 microelectrodes (Revsbech, 1989) connected to
a pA-meter (PA2000 or Microsensor Multimeter, Unisense,
November 2012 | Volume 3 | Article 402 | 13
Kühl et al.
Denmark). The O2 microsensors exhibited a fast response time
(t 90 < 0.5 s), low stirring sensitivity (<1–2%) and had tip diameters of 25–50 µm. In L. patella, it was necessary to make an incision
with a thin (29G) hypodermic needle prior to measurements. The
O2 microsensors were linearly calibrated from sensor readings
in O2 -free solution (seawater amended with sodium dithionite)
and in aerated seawater at experimental temperature and salinity
(100% atmospheric saturation).
Microscale pH measurements were attempted with pH glass
needle electrodes (pH-N, Unisense AS, Aarhus, Denmark; Kühl
and Revsbech, 2001) connected to a high impedance mV meter
(Keithley, USA) and calibrated at experimental temperature in
standard pH buffers at pH 4, 7, and 10. The sensors exhibited a
near-ideal Nernstian response of 55–59 mV per pH unit change.
However, the pH sensors were too fragile for profiling pH in the
sturdy tunic of L. patella and we observed apparent local release
of acid vacuoles, when inserting the relatively large needle sensors,
that changed the pH microenvironment dramatically down to pH
2–3 within seconds before slowly rising again to ambient pH over
some minutes.
For microprofiling, the microsensors were mounted in a PCcontrolled motorized micromanipulator system (Unisense, Denmark) that enabled automated profiling (in vertical steps of
50–100 µm) and data acquisition via dedicated software (Profix,
Pyroscience, Germany).
IMAGING OF O2 AND PH DISTRIBUTION AND DYNAMICS
The spatial distribution and dynamics of O2 concentration and
pH in L. patella was imaged on a vertical cut specimen, which was
mounted tightly up against the transparent flow chamber wall
with help of two hypodermic needles fixed in the soft chamber bottom. The planar optodes (see below) were positioned
between the specimen and the inner wall of the flow chamber. The temperature in the flow chamber was kept constant at
26˚C. Actinic light was provided vertically from above using a
fiber-optic halogen lamp (LG-PS2, Olympus, Japan). To avoid
interference, the lamp was briefly switched off during the image
acquisition. The irradiance of the actinic light at the level of the
ascidian test surface was determined in water with a scalar irradiance mini sensor (Model US-SQS/L, Walz GmbH, Effeltrich,
Germany) coupled to a calibrated irradiance meter (ULM-500,
Walz GmbH, Effeltrich, Germany). Photographic images of the
vertically cut ascidian surface mounted against the transparent
flow chamber were taken with a digital camera (Canon 5D, Canon,
Japan).
Sensor materials
The fundamentals of sensor fabrication and solute imaging with
planar optodes as well as details of sensor synthesis and physicochemical properties are described in detail elsewhere (e.g., Borisov
et al., 2008, 2009a,b; Kühl and Polerecky, 2008; Kühl et al.,
2007b, 2008; Mayr et al., 2009; Larsen et al., 2011; Staal et al.,
2011; Fabricius-Dyg et al., 2012). Planar optodes for O2 imaging
were prepared by first dissolving 3 mg of platinum(II) tetra(4fluoro)phenyltetrabenzoporphyrin (=PtTPTBPF; Borisov et al.,
2008, 2009a) and 200 mg of polystyrene (MW 250,000, Aldrich,
USA) in 1.8 g of chloroform. This “cocktail” was knife-coated onto
Frontiers in Microbiology | Aquatic Microbiology
Prochloron microenvironment
a transparent poly(ethylene terephthalate) support foil (Mylar,
Goodfellow, USA). The thickness of the sensing layer after evaporation of the solvent was ∼2.5 µm. The O2 sensor chemistry
possesses exceptional brightness due to high molar absorption
coefficients and luminescence quantum yields of the indicators
and enables excitation with red light (λmax 615 nm). The NIR
emission at ∼765 nm allows minimization of scattering effects
and interfering chlorophyll fluorescence.
Planar optodes for pH imaging were prepared by first dissolving 3 mg of the coumarin dye Macrolex®Fluorescent Yellow (=MY;
Simon and Werner GmbH)1 , 3 mg of lipophilic 1-hydroxypyrene3,6,8-tris-bis(2-ethylhexyl)sulfonamide [=HPTS(DHA)3 ; Borisov
et al., 2009b] and 200 mg of Hydrogel D4 (Cardiotech)2 in
1.8 g of ethanol:water (9:1 v/v). The pH sensor “cocktail” was
also coated onto a transparent Mylar support foil. The thickness of the pH sensing layer after evaporation of the solvent was
∼7.5 µm.
Since the absorption spectrum of the basic form of the pH
indicator perfectly matches the emission spectrum of the MY
reference dye, Förster Resonance Energy Transfer (FRET) leads
to the emission from HPTS(DHA)3 at high pH (Figure 9).
The protonated form of HPTS(DHA)3 absorbs at about 450 nm,
therefore, no FRET is observed at lower pH and the emission from MY is maximal. Thus, the system allows referenced ratiometric imaging of pH by monitoring the emissions
from the two dyes – MY at ∼505 nm and HPTS(DHA)3 at
∼555 nm.
Imaging of O2
Planar O2 optodes were read out with a lifetime imaging system [see details in Holst et al., 1998) employing a monochrome
gated CCD camera (Sensicam-Sensimod, PCO, Kehlheim, Germany)] equipped with a Schneider-Kreuznach Xenoplan 1.4/23
CCTV-Lens and a R-720 long-pass filter (Edmunds Optics, USA)
mounted in front of the lens. Two red-orange high power LEDs
(617 nm, 1 W, Luxeon, Philips Lumileds, USA) were used for excitation, as controlled by a custom-built PC-controlled pulse-delay
generator. A rapid lifetime determination method was used to
determine the O2 -dependent luminescence decay time τ by measuring luminescence intensities in two time windows (1–41 and
26–66 µs, respectively) after the excitation pulse:
τ=
t1 − t2
,
ln I1 I2
(1)
where t 1 and t 2 represent the time corresponding to the start of
the first and the second window, respectively (1 and 26 µs in our
setup), and I 1 and I 2 are the luminescence intensities acquired in
the respective time windows.
A two point calibration (in air saturated and anoxic seawater,
respectively) was used to calibrate the τ vs. O2 oxygen response.
A 2% w/w aqueous solution of sodium dithionite was used
for deoxygenation. The following equation was used to obtain
1 http://www.simon-und-werner.de
2 http://www.cardiotech-inc.com
November 2012 | Volume 3 | Article 402 | 14
Kühl et al.
Prochloron microenvironment
the calibration curve and to convert the measured decay times
into pO2 :
τ
f
=
+1−f,
τ0
1 + KSV [O2 ]
(2)
where f = 0.91 (determined from the calibration for the
PtTPTBPF/PS optodes obtained in the frequency domain measurements), τ0 is the decay time in the anoxic solution
(53.2 µs), and K SV is the Stern–Volmer constant (0.0196 hPa−1 )
as determined from the decay times at air saturation and under
anoxic conditions under experimental temperature and salinity.
We used Matlab 7.10 (Mathworks Inc., USA) to calculate
pseudocolor decay time images from the original data and,
subsequently pseudocolor O2 concentration images.
Imaging of pH
The pH optodes were read out with a new spectral camera system
(Spectrocam, Ocean Thin Films, USA) consisting of a CCD camera
combined with a fast-switching filter wheel (eight filter positions) and equipped with a Distagon TX2.8/25 mm ZF lens (Carl
Zeiss AG, Germany). A long-pass glass filter (OG 490, SchneiderKreuznach GmbH, Germany) was mounted in front of the lens to
eliminate interference from the reflected excitation light. The pH
optode was excited with blue light (λmax 460 nm, as determined
with a fiber-optic spectrometer, USB2000+, Ocean Optics, USA)
from a torch (Bluestar, NightSea, Bedford, MA, USA). The emission of the pH sensor was monitored in two spectral windows:
“window 1” (filter position with a Spectrocam 475/100 nm band
width interference filter) and “window 2” (filter position with a
Spectrocam 550/100 nm band width interference filter).
The emission of HPTS(DHA)3 was imaged with the
550/100 nm filter. Notably, some emission from MY is also
detected in this window (Figure 9A). The emission from MY was
detected using the 475/100 nm bandpass filter. Thus, the effective
window for monitoring the green fluorescence of MY was 490–
525 nm. Subsequently, pH was evaluated by the ratio of the two
windows: R = (window 2/window 1) × 1200.
The pH optode was calibrated by ratiometric imaging in
seawater at experimental temperature and salinity, wherein specific pH values were adjusted by addition of 0.1 M HCL or
1 M NaOH. The seawater pH at different calibration points
was measured with a calibrated pH glass electrode connected
to a pH meter (UB-10, Denver Instruments, USA). Calibration curves were obtained both for a fresh sensor foil and a
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AUTHOR CONTRIBUTIONS
Michael Kühl, Lars Behrendt, Sergey M. Borisov, Ulrich
Schreiber, and Anthony W. D. Larkum designed and performed
research; Michael Kühl, Erik Trampe, Klaus Qvortrup, Ulrich
Schreiber, Sergey M. Borisov, and Ingo Klimant contributed new
reagents/analytic tools; Michael Kühl, and Sergey M. Borisov analyzed data; and Michael Kühl, Lars Behrendt, Sergey M. Borisov,
and Anthony W. D. Larkum wrote the paper with editorial help
from all co-authors.
ACKNOWLEDGMENTS
This study was supported by grants from the Danish Natural Science Research Council, the Carlsberg Foundation (Denmark), and
the Australian Research Council. We acknowledge the excellent
technical support by the staff at Heron Island Research Station,
University of Queensland. We thank Anni Glud and Lars Rickelt for
construction of microsensors, Roland Thar for developing the customized data acquisition software used in this study, and Murray
Badger for helpful advice on possible inorganic carbon transport
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(CFIM), Faculty of Health and Medical Sciences, University of
Copenhagen is acknowledged for access to electron microscopes.
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Conflict of Interest Statement: The
authors declare that the research was
conducted in the absence of any
commercial or financial relationships
that could be construed as a potential
conflict of interest.
Received: 28 August 2012; paper pending published: 14 October 2012; accepted:
02 November 2012; published online: 30
November 2012.
Citation: Kühl M, Behrendt L, Trampe
E, Qvortrup K, Schreiber U, Borisov
SM, Klimant I and Larkum AWD
(2012) Microenvironmental ecology
of the chlorophyll b-containing symbiotic cyanobacterium Prochloron in
the didemnid ascidian Lissoclinum
patella. Front. Microbio. 3:402. doi:
10.3389/fmicb.2012.00402
This article was submitted to Frontiers
in Aquatic Microbiology, a specialty of
Frontiers in Microbiology.
Copyright © 2012 Kühl, Behrendt ,
Trampe, Qvortrup, Schreiber, Borisov,
Klimant and Larkum. This is an openaccess article distributed under the terms
of the Creative Commons Attribution
License, which permits use, distribution
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