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RESEARCH LETTER
Coral mucus-associated bacterial communities from natural and
aquarium environments
Netta Kooperman1, Eitan Ben-Dov1,2, Esti Kramarsky-Winter3, Zeev Barak4 & Ariel Kushmaro1
1
Department of Biotechnology Engineering, Ben-Gurion University of the Negev, Be’er Sheva, Israel; 2Achva Academic College, MP Shikmim, Israel;
Department of Zoology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel; and 4Department of Life Sciences, Ben-Gurion
University of the Negev, Be’er Sheva, Israel
3
Correspondence: Ariel Kushmaro,
Department of Biotechnology Engineering,
Ben-Gurion University of the Negev, P. O. Box
653, Be’er Sheva, 84105, Israel. Tel.: 1972 8
6479 024; fax: 1972 8 6472 983; e-mail:
arielkus@ bgu.ac.il
Received 15 May 2007; revised 23 July 2007;
accepted 10 August 2007.
First published online October 2007.
DOI:10.1111/j.1574-6968.2007.00921.x
Editor: Herman Bothe
Keywords
coral; Fungia granulosa ; mucus microbiota;
Red Sea.
Abstract
The microbial biota dwelling in the mucus, on the surface, and in the tissues of
many coral species may have an important role in holobiont physiology and
health. This microbiota differs with coral species, water depth, and geographic
location. Here we compare the surface mucus microbiota of the coral Fungia
granulosa from the natural environment with that from individuals maintained in
aquaria. Molecular analysis revealed that the microbial community of the mucus
microlayer of the coral F. granulosa includes a wide range of bacteria and that these
change with environment. Coral mucus from the natural environment contained a
significantly higher diversity of microorganisms than did mucus from corals
maintained in the closed-system aquaria. A microbial community shift, with the
loss of several groups, including actinobacterial and cyanobacterial groups, was
observed in corals maintained in aquaria. The most abundant bacterial class in
F. granulosa mucus was the Alphaproteobacteria, regardless of whether the corals
were from aquaria or freshly collected from their natural environment. A
significantly higher percentage of bacteria from the Betaproteobacteria class was
evident in aquarium corals (24%) when compared with corals from the natural
environment (3%). The differences in mucus-inhabiting microbial communities
between corals from captive and natural environments suggest an adaptation of
the mucus bacterial communities to the different conditions.
Introduction
Coral reefs are diverse and important communities in
tropical and subtropical marine environments. Hermatypic
corals play a key role in forming the structure of coral reefs
and in providing substrata and shelter for a wide variety of
organisms. Studies have revealed a dynamic microbial biota
living in the mucus, on the surface, and in the tissues of
many coral species (Ritchie & Smith, 1997; Rohwer et al.,
2001; Guppy & Bythell, 2006; Koren & Rosenberg, 2006).
Coral microorganisms may be mutualistic or pathogenic, or
they may provide other important functions in the ecosystem (Kushmaro et al., 1996; Santavy & Peters, 1997; Harvell
et al., 1999; Ben-Haim et al., 2003). One possible function of
microorganisms found on coral holobiont surfaces may be
to provide the corals with protection from pathogens by
means of interspecific competition and/or secretion of
antibiotic substances (Rohwer et al., 2002; Reshef et al.,
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c
2006; Ritchie, 2006; Rosenberg et al., 2007). Another role for
these microorganisms may be to supply the coral with
nitrogen and phosphorous, which are not provided by their
symbiotic zooxanthellae (Sorokin, 1973a, b, 1978; Anthony,
1999, 2000; Rosenfeld et al., 1999; Anthony & Fabricius,
2000).
The coral surface microlayer is highly productive, mucusrich, and extends a few millimetres above the surface tissue
of the coral (Paul et al., 1986; Brown & Bythell, 2005).
Specialized mucus cells present in the coral epidermis
secrete the mucus layer, which contains polymers that form
a highly hydrated viscoelastic polymeric gel, consisting of
fucose, arabinose, mannose, galactose and small amounts of
glucose residues (Meikle et al., 1988). Ducklow & Mitchell
(1979) reported that coral mucus sustains high bacterial
growth, presumably resulting from the degradation of the
mucus constituents. They furthermore reported that mucus,
FEMS Microbiol Lett 276 (2007) 106–113
107
Coral mucus-associated bacterial communities
its degradation products, and the bacteria living on this
mucus may be used as nutrient sources by other organisms.
In addition, coral physiological status and environmental
parameters such as water motion, irradiation and availability of nutrients play a role in the stability and composition of
the mucus layer (Brown & Bythell, 2005). It is therefore
likely that environmental (e.g. temperature, irradiance,
nutrient availability) conditions together with the coral’s
physiological condition (e.g. health, reproductive status,
etc.) determine the microbial community associated with a
coral holobiont. In turn, changes in microbial communities
may affect coral physiology. In this study we assess the
mucus microbial communities of the coral Fungia granulosa
from natural and aquarium environments.
Materials and methods
Sample collection and laboratory maintenance
Fungia granulosa individuals of similar sizes (5 cm in
diameter) were collected from the area where they are most
abundant, near the Inter-University Institute for Marine
Science in the Gulf of Eilat, Red Sea (29151 0 N, 34194 0 E)
(Kramarsky-Winter & Loya, 1998) at depths between 9 and
24 m, during the early summer of 2004. Eight individual
corals were maintained in separate aerated 2-L aquaria with
artificial seawater (Instant Ocean) under a controlled environment at a constant temperature of 22 1C and with a light
regime of 12 : 12 h (light : dark). The water was changed
every three days. After an acclimation period of three weeks,
mucus for molecular analysis was collected by rubbing the
coral surface with bacteriological loops. Samples collected
from the coral mucus of aquarium corals are termed CMAC.
Coral mucus from the natural environment (CMNE) of
nine F. granulosa individuals was sampled in situ during the
same period and at the same site. To do this, sterile
bacteriological loops (three for each sample) were inserted
into dry, sterile 15-mL polypropylene centrifuge tubes. The
tubes were opened upside down underwater to prevent the
entrance of seawater; the loops were then extricated and
rubbed on the coral surface. Compressed air was added to
the tubes to remove seawater, and the loops were reinserted
into the tubes. The tubes were then sealed in the inverted
position, and once brought to the surface were immediately
placed on ice.
DNA extraction
Genomic DNA from the mucus microlayer was extracted
using a NucleoSpin food purification kit (Macherey-Nagel,
Düren, Germany). Zircon silica beads 0.5 mm in diameter
were used rather than the homogenization of samples
recommended in the kit instructions. Genomic DNA was
FEMS Microbiol Lett 276 (2007) 106–113
eluted using 50 mL of elution buffer and stored at 20 1C. To
extract genomic DNA from the water column, 2 L of seawater
collected from the same depths was filtered through a 0.2-mm
filter, and DNA was extracted as described above.
PCR amplification
Total DNA was amplified with a Mastercycler gradient
thermocycler (Eppendorf, Westbury, NY) by PCR using
specific 16S rRNA primers for bacteria. Primers used for the
construction of clone libraries were: forward primer
8F (5 0 -GGATCCAGACTTTGAT(C/T)(A/C)TGGCTCAG),
taken and modified from Felske et al. (1997) (the 8F primer
was shortened from the 5 0 end); and reverse primer 907R
(5 0 -CCGTCAATTCCTTT(A/G)AGTTT-3 0 ), taken from
Muyzer et al. (1996). Reaction mixtures consisted of 12.5 mL
of Reddy-Mix (PCR Master mix containing 1.5 mM MgCl2
and 0.2 mM concentration of each deoxynucleoside triphosphate) (ABgene, Surrey, UK), 1 pmol of each of the forward
and reverse primers, 1 to 2 mL of the sample preparation, and
water to bring the total volume to 25 mL. An initial denaturation hot start of 4 min at 95 1C was followed by 30 cycles of
the following incubation pattern: 94 1C for 30 s, 53 to 56 1C
for 40 s, and 72 1C for 105 s. The procedure was completed
with a final elongation step at 72 1C for 20 min.
Clone library construction and sequencing
PCR products were purified by electrophoresis through a
0.8% agarose gel (Sigma), stained with ethidium bromide,
and visualized on a UV transilluminator. The c. 900-bp
heterologous rRNA gene products were excised from the gel,
and the DNA was purified from the gel slice using the
Wizard SV gel and PCR clean-up system (Promega, Madison, WI). The gel-purified PCR products were cloned into
the pGEM-T Easy vector (Promega) and transformed into
calcium chloride-competent XL MRF’ Escherichia coli cells
according to the manufacturer’s instructions and standard
techniques (Sambrook & Russell, 2001). Plasmid DNA was
isolated from individual clones using the Wizard Plus SV
Minipreps DNA purification system (Promega).
Aliquots from a subset of the samples of purified plasmid
DNA were digested with the restriction enzyme EcoRI (MBI
Fermentas) for more than 4 h at 37 1C, and the digested
product was separated by electrophoresis on a 1% agarose
gel (agarose low electroendosmosis; Hispanagar, Spain).
After being staining with ethidium bromide, the bands were
visualized on a UV transilluminator to select clones containing the appropriately sized insert. Sequencing with 8F and
341F primers was performed with an ABI PRISM dye
terminator cycle sequencing ready reaction kit with AmpliTaq DNA polymerase FS (Taq-FS, a member of the Taq
F667Y family) and a DNA sequencer ABI model 373A
system (Perkin-Elmer).
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108
Sequence analyses
All the rRNA gene sequences of each group were first
compared with those in the GenBank database using the
BLAST network service (http://www.ncbi.nlm.nih.gov/blast/
blast.cgi). CLASSIFIER (version 1.0; assign 16S rRNA sequences
to a taxonomical hierarchy) and LIBRARY COMPARE [compare
two sequence libraries using the Ribosomal Database Project
(RDP) CLASSIFIER] available at the Ribosomal Database
Project-II
website
(http://rdp.cme.msu.edu/index.jsp,
Maidak et al., 1999) were used to find diversity on different
ranks (phylum, class, order etc.) of related sequences. To
control for the occurrence of possibly chimeric sequences,
all sequenced clones were analysed with the CHIMERA CHECK
program of the RDP database (version 2.7; Maidak et al.,
1999). The sequences from appropriate libraries
were aligned using CLUSTALW with the MEGA package
(Kumar et al., 2004), and positions not sequenced in all
isolates or with alignment uncertainties were removed.
Phylogenetic trees were constructed with the neighbourjoining method (Saito & Nei, 1987) using the MEGA package
(Kumar et al., 2004). Bootstrap resampling analysis (Felsenstein, 1985) for 100 replicates was performed to estimate the
confidence of tree topologies.
Estimation of community richness
N. Kooperman et al.
were retrieved using clone libraries. In addition, 22
sequences were retrieved from the surrounding water using
clone libraries. The diversity and distribution of the 16S
rRNA gene sequences denoting the various bacterial groups
from seawater samples were very different from those found
in the pooled coral mucus samples (Fig. 1). In the seawater
samples (Fig. 1a), one of the most abundant groups, at 30%
of the sequences, was the cyanobacteria, whereas the corresponding abundance was 9% in the CMNE (Fig. 1b). The
Gammaproteobacteria group accounted for only 5% of the
sequences in seawater as opposed to 17% in the CMNE (Fig.
1a and b). There were a few ribotypes with similarities of
(a)
Alphaproteobacteria
35%
Cyanobacteria
30%
CFB
10%
Betaproteobacteria
5%
Gammaproteobacteria
5%
(b)
Cyanobacteria
9%
Operational taxonomic units (OTUs) for the purposes of
community analysis were defined by a 3% (cut-off 97%, for
species-level similarity) and 17% (cut-off 83%, for phylumclass-level similarity) difference in nucleic acid sequences, as
determined using the furthest neighbor algorithm in DOTUR
(Schloss & Handelsman, 2005). Rarefaction, richness, and
diversity statistics were also calculated using DOTUR, including the nonparametric richness estimators Chao1 and the
Shannon diversity index.
Nucleotide sequence accession numbers
Unidentified
15%
Unidentified
12%
Firmicutes
4%
Alphaproteobacteria
33%
Gamma
proteobacteria
17%
Beta
proteobacteria
3%
CFB
9% Actinobacteria
7%
(c)
The sequences from this study are available through GenBank under accession numbers DQ117312DQ117435
(coral mucus), and DQ417904DQ417925 (seawater).
Planctomycetes
3%
Verrucomicrobia
3%
Unidentified
7%
Alphaproteobacteria
33%
Gammaproteobacteria
30%
Results
Molecular analysis revealed that the microbial community
of the mucus microlayer of the coral F. granulosa includes a
wide range of bacteria and that these change with environment. The microbial groups found on the coral mucus range
from obligatory aerobes to anaerobic bacteria and photosynthetic bacteria, and cyanobacteria that are known to be
nitrogen fixers.
Mucus was collected from nine coral individuals from the
natural environment and from eight individuals maintained
in aquaria, from which 76 and 48 sequences, respectively,
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CFB
6%
Betaproteobacteria
24%
Fig. 1. Distribution of bacterial 16S rRNA gene sequences from clone
libraries. (a) seawater, (b) mucus of Fungia granulosa from the Red Sea,
and (c) mucus of F. granulosa kept in aquaria. A 97100% match of the
unknown clone with the GenBank dataset was considered an accurate
identification to the species level. Sequences with similarities of 90% or
less were considered unidentified prokaryotes.
FEMS Microbiol Lett 276 (2007) 106–113
109
Coral mucus-associated bacterial communities
Fig. 2. Phylogenetic tree based on 16S rRNA gene sequences that were retrieved from mucus of Fungia granulosa from the Red Sea (RS.Muc) and
mucus of F. granulosa kept in aquaria (Aq.Muc). The tree was constructed using the neighbour-joining method (Saito & Nei, 1987) with the MEGA
package (Kumar et al., 2004) using partial sequences of 16S rRNA genes. The bar represents five substitutions per 100 nucleotide positions. Bootstrap
values (Felsenstein, 1985) are indicated at branch nodes.
FEMS Microbiol Lett 276 (2007) 106–113
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c
110
N. Kooperman et al.
97% and above that appeared both in the seawater and in
the mucus communities.
Overall, significant differences (2w d.f. = 1, P o 0.05)
were observed in mucus-associated microorganisms in
corals from the natural and the aquarium environments.
The Alphaproteobacteria group was the most abundant
bacterial class from mucus of F. granulose, regardless of
whether the corals were kept in aquaria (Fig. 1c) or collected
from the environment (Fig. 1b). In addition, members of
the Gammaproteobacteria and Cytophagales-FlavobacteriaBacteroidetes (CFB) were abundant in all of the corals.
Corals maintained in closed aquarium systems, using artificial seawater, showed a significantly lower diversity of
bacteria than did corals from the environment (Fig. 1b and
c). Aquarium-acclimated individuals were lacking several
bacterial groups, including actinobacteria and cyanobacteria, that were found in CMNE. Moreover, there was a distinct
and significantly higher percentage (2w d.f. = 1, P o 0.05) of
bacteria from the Betaproteobacteria class in CMAC (24%)
than in CMNE (3%). This class was mainly composed of the
Burkholderiales group. In this group, three sequences,
one from CMNE (RS.Muc.122) and two from CMAC
(Aq.Muc.099 and Aq.Muc.098), showed above 99% identity
(Fig. 2b). The Gammaproteobacteria group also accounted
for a higher percentage of the microbial community
in CMAC than in CMNE, but this difference was not
significant.
A phylogenetic tree of 16S rRNA gene sequences from
coral mucus from the natural environment and from the
aquaria is presented in Fig. 2. The phylogenetic tree adds
information at a higher resolution than in Fig. 1 and
provides information at the levels of genus and specific
clones. At the genus level, sequences retrieved from the
mucus layer belong to Janthinobacterium, Alcaligenes (Betaproteobacteria), Pseudomonas, Stenotrophomonas, Shewanella,
Alteromonas, Pseudoalteromonas (Gammaproteobacteria),
Rhizobium and Rhodobacteraceae (Alphaproteobacteria).
Rarefaction analysis (for the first 43 sequences) at the
species level (cut-off 97%) and at the phylum-class level
(cut-off 83%) revealed 29 and 19 OTUs for the mucus of
corals from the natural environment, and 30 and 15 OTUs
for the mucus of corals from aquarium samples (Table 1),
respectively. However, according Chao1 richness estimator at
the species and at the phylum-class levels revealed 88 and
22 OTUs for the mucus of corals from the Red Sea and 72
and 36 OTUs for the mucus of corals from aquarium
samples, respectively, and the Shannon–Weaver diversity
index was 3.45 and 2.83 for the natural environment samples
and 3.2 and 2.42 for the aquarium samples (Table 1).
Discussion
This study provides a glimpse into the mucus-associated
microbial communities of the coral F. granulosa in the
natural environment and in aquaria. Our results present an
overall high diversity of bacteria in the mucus microlayer of
this coral. Over 50% of the sequences were putative novel
species (i.e. less than 97% similarity to GenBank entries),
and 20% were putative novel genera (i.e. less than 93%
similarity to GenBank entries). In general, we were unable
to find specific associations of bacterial ribotypes with
F. granulosa from either environment. This may have been
because of the high diversity of coral mucus-associated
microorganisms. One exception was a recurrent ribotype
with 97% similarity to Ochrobactrum, a genus that belongs
to the Alphaproteobacteria. This ribotype was found repeatedly in the mucus of individual F. granulosa from the natural
environment. Interestingly, this genus is prevalent in other
coral species (Rohwer et al., 2002), and is known to be
present in marine biofilms (Lee et al., 2003). Furthermore,
preliminary results (Kooperman, 2005) have shown that
there are three groups belonging to the Archaea of repeatedly occurring ribotypes in mucus collected from F. granulosa individuals at different times and from different
locations.
Some overlap was found between the coral microbiota
and that of the surrounding seawater, indicating a water mucus interaction. There was a dominance of the Alphaproteobacteria in the natural seawater in Eilat (Fig. 1a), and
this was the most abundant bacterial group on the coral
mucus surface, regardless of environment (Fig. 1b). There
have been similar findings from a number of studies from
Table 1. Observed and estimated richnesses of 16S rRNA gene libraries from mucus of Fungia granulosa samples in aquaria and from the Red Sea,
as estimated by rarefaction analysis, the Shannon–Weaver diversity index, and the Chao1 richness estimator computed using DOTUR
No. of OTUs
Mucus clone library
CMAC
CMNE
w
Chao1 value
No. of clones sequenced Cut-off 97% Cut-off 83% Cut-off 97%
43
65
30
38 (29)z
15
21 (19)z
Shannon–Weaver index
Cut-off 83%
Cut-off 97%
Cut-off 83%
72 (45 152) 36 (20 101) 3.2 (2.93 3.48) 2.42 (2.16 2.67)
88 (56 180) 22 (21 31) 3.45 (3.24 3.65) 2.83 (2.65 3.01)
Numbers of OTUs, the Chao1 estimated richness and the Shannon–Weaver diversity index are shown for both 3% and 17% differences in nucleic acid
sequence alignments. Numbers in parentheses are lower and upper 95% confidence intervals.
w
CMAC: coral mucus from aquarium corals; CMNE: coral mucus from the natural environment.
z
The numbers in parentheses refer to the numbers of OTUs obtained for the first 43 sequences.
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FEMS Microbiol Lett 276 (2007) 106–113
111
Coral mucus-associated bacterial communities
various geographic locations, including the Caribbean and
the Great Barrier Reef (Rohwer et al., 2001, 2002; Bourne &
Munn, 2005). This dominance resembles that found in the
coral reef-associated water column from a variety of geographical locations (Rappe et al., 2000; Bourne & Munn,
2005). Interestingly, in contrast to these results, Frias-Lopez
et al. (2002) found that the Gammaproteobacteria was the
most abundant group in the coral Montastrea cavernosa
from the Caribbean. These differences may be the result of
host-species differences as well as of the fact that these
researchers analysed the microbial community of a mixture
of coral mucus, tissues and skeleton, and not only of the
coral mucus. Because the coral mucus layer is in constant
association with the surrounding water column, and bacteria may shift from the water column to the mucus and vice
versa, it is not surprising that similar ribotypes in the mucus
and seawater samples were evident. Approximately 7% of
the microbial communities found on the surface of
F. granulosa from the natural environment were Actinobacteria (Fig. 1b). Similarly, in a recent publication Lampert
et al. (2006) found that 23% of 22 isolates from the mucus of
another fungiid species, Fungia scutaria, collected from its
ambient environment belonged to the Actinobacteria.
The microbial community diversity found in mucus from
natural F. granulosa individuals is similar to that of the
coral-associated community of other naturally occurring
corals (Rowher et al., 2002; Bourne & Munn, 2005) from
other environments. Overall, there was a lower microbial
community diversity in aquarium-coral mucus (Fig. 1c)
than in mucus from corals from the natural environment
(Fig. 1b). Rarefaction analysis (number of OTUs) and the
Shannon–Weaver diversity index (3.45 and 2.83 for CMNE
samples, and 3.2 and 2.42 for CMAC samples for cut-offs of
97% and 83%, respectively) (Table 1) both support this
observation. It is noteworthy that the Shannon–Weaver
diversity index (3.2) obtained from mucus of the coral
Pocillopora damicornis located on the Great Barrier Reef
(Bourne & Munn, 2005) is similar to the diversity index
obtained from corals in this study. The lower microbial
diversity in aquarium-coral mucus was characterized by the
absence of several groups and by higher proportions of
others. The shift in microbial community equilibrium, as
observed, for example, in the greater proportions of Betaproteobacteria in aquarium corals (Fig. 1c), may either be the
result of or be part of the cause of overall physiological
changes in the holobiont. For example, the loss of some of
the Actinobacteria, known for their production of many
bioactive compounds (Magarvey et al., 2004; Fiedler et al.,
2005; Jensen et al., 2005), may affect the susceptibility of
these corals to pathogens (e.g. Rohwer et al., 2002). The
identification of groups such as the Burkholderiales group
(Betaproteobacteria) (Fig. 2a and b), which includes a
number of pathogenic species, particularly some that cause
FEMS Microbiol Lett 276 (2007) 106–113
plant diseases (Burkholder, 1950; Lincoln et al., 1999), in
CMAC may also represent such a shift. Interestingly, CMAC
bacterial sequences Aq.Muc.098 and Aq.Muc.099 and sequence RS.Muc.122 from CMNE showed 99% similarity to
Alcaligenes faecalis strain BC2001 (accession no. AY667065),
a bacterium with anti-nematode activity (Zhou & Zheng,
pers. commun.). This activity may be important for a coral
that resides on and in close contact to the substrate. In
addition, CMAC contained a number of sequences (e.g.
Aq.Muc.090, Aq.Muc.094 and Aq.Muc.041, Aq.Muc.044)
belonging to the genera Pseudomonas and Alteromonas (Fig.
2c and f, respectively) that were not apparent in CMNE.
Members of these genera are known to be associated with
diseased corals (Frias-Lopez et al., 2002), and in this study
were detected only in aquarium corals. By contrast, we
found
members
(RS.Muc.150,
RS.Muc.153
and
RS.Muc.195; Fig. 2e) of the genus Shewanella, a group
known to be associated with normal coral flora (Rohwer
et al., 2001), only in corals from the marine environment.
These results indicate that the physiology of aquarium corals
may be different from that of corals from the natural
environment, which in turn influences the microbial community structure.
Studies have shown that coral-associated microbial communities can change as a function of depth, water quality,
geographic location, and colony health (Rohwer et al., 2001,
2002; Frias-Lopez et al., 2002; Kooperman, 2005; Reshef
et al., 2006; Klaus et al., 2007; Rosenberg et al., 2007). These
changes are likely to accompany changes in coral physiological function. Indeed, Reshef et al. (2006) suggested that
corals could adapt rapidly to changing environmental conditions by altering their population of symbiotic bacteria.
They further posited that a dynamic relationship exists
between symbiotic microorganisms and environmental conditions that brings about the selection of the most advantageous coral holobiont in changing environmental
conditions. On the other hand, the recurrence of microorganisms in corals from different environments or physiological states may indicate an obligate symbiont. The
recurrence of sequences from the Betaproteobateria found
in mucus from corals from the natural environment
(RS.Muc.122) and from the aquarium (Aq.Muc.099 and
Aq.Muc.098) with above 99% identity (see Fig. 2b) may
therefore indicate an obligate symbiont. This group merits
further study to attempt to assess its relationship to its coral
host species.
Similar to what was found in aquarium corals, in diseased
corals there was a reduction in microbial group numbers
compared with flora from healthy colonies (Pantos et al.,
2003). If these changes are caused by bacterial groups being
selected for as a result of environmental factors acting
directly on the bacterial communities, or indirectly by
means of changes in the coral physiological response (Klaus
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c
112
et al., 2005), this may provide us with clues regarding their
susceptibility to disease. The differences in microbial communities in mucus between corals from captive and natural
environments suggest an adaptation of the mucus bacterial
communities to the various conditions that may affect the
coral holobiont physiology.
Acknowledgements
This work was supported by ISF grant no. 511/02-1. We
would like to thank the H. Steinitz Marine Biological
Laboratory at Eilat for use of their facilities. We thank
Nachshon Siboni, Michal Lidor and Orr Shapiro for sample
collection and for technical support.
References
Anthony KRN (1999) Coral suspension feeding on fine
particulate matter. J Exp Mar Biol Ecol 232: 85–106.
Anthony KRN (2000) Enhanced particle-feeding capacity of
corals on turbid reefs (Great Barrier Reef, Australia). Coral
Reefs 19: 59–67.
Anthony KRN & Fabricius EF (2000) Shifting roles of
heterotrophy and autotrophy in coral energetics under
varying turbidity. J Exp Mar Biol Ecol 252: 221–253.
Ben-Haim Y, Thompson FL, Thompson CC, Cnockaert MC,
Hoste B, Swings J & Rosenberg E (2003) Vibrio coralliilyticus
sp. nov., a temperature-dependent pathogen of the coral
Pocillopora damicornis. Int J Syst Evol Microbiol 53: 309–315.
Bourne DG & Munn CB (2005) Diversity of bacteria associated
with the coral Pocillopora damicornis from the Great Barrier
Reef. Environ Microbiol 7: 1162–1174.
Brown BE & Bythell JC (2005) Perspectives on mucus secretion
in reef corals. Mar Ecol Prog Ser 296: 291–309.
Burkholder WH (1950) Sour skin: a bacterial rot of onion bulbs.
Phytopathology 40: 115–117.
Ducklow WH & Mitchell R (1979) Bacterial populations and
adaptations in the mucus layers on living corals. Limnol
Oceanog 24: 715–725.
Felsenstein J (1985) Confidence limits of phylogenies: an
approach using the bootstrap. Evolution 39: 783–791.
Felske A, Rheims H, Wolterink A, Stackebrandt E & Akkermans
ADL (1997) Ribosome analysis reveals prominent activity of
an uncultured member of the class Actinobacteria in grassland
soils. Microbiology 143: 2983–2989.
Fiedler HP, Bruntner C, Bull AT, Ward AC, Goodfellow M,
Potterat O, Puder C & Mihm G (2005) Marine actinomycetes
as a source of novel secondary metabolites. Antonie van
Leeuwenhoek 87: 37–42.
Frias-Lopez J, Zerkle AL, Bonheyo GT & Fouke BW (2002)
Partitioning of bacterial communities between seawater and
healthy, black band diseased and dead coral surfaces. Appl
Environ Microbiol 68: 2214–2228.
2007 Federation of European Microbiological Societies
Published by Blackwell Publishing Ltd. All rights reserved
c
N. Kooperman et al.
Guppy R & Bythell JC (2006) Environmental effects on
bacterial diversity in the surface mucus layer of the reef
coral Montastraea faveolata. Mar Ecol Prog Ser 328: 133–142.
Harvell CD, Kim K, Burkholder JM et al. (1999) Review: marine
ecology-emerging marine diseases–climate links and
anthropogenic factors. Science 285: 1505–1510.
Jensen PR, Mincer TJ, Williams PG & Fenical W (2005) Marine
actinomycete diversity and natural product discovery. Antonie
van Leeuwenhoek 87: 43–48.
Klaus JS, Frias-Lopez J, Bonheyo GT, Heikoop JM & Fouke BW
(2005) Bacterial communities inhabiting the healthy tissues of
two Caribbean reef corals: interspecific and spatial variation.
Coral Reefs 24: 129–137.
Klaus JS, Janse I, Heikoop JM, Sanford RA & Fouke BW (2007)
Coral microbial communities, zooxanthellae and mucus along
gradients of seawater depth and coastal pollution. Environ
Microbiol 9: 1291–1305.
Kooperman N (2005) Microbial diversity of Fungia granulosa
mucus layer. MSc Thesis, Ben-Gurion University, Israel
Koren O & Rosenberg E (2006) Bacteria associated with mucus
and tissues of the coral Oculina patagonica in summer and
winter. Appl Environ Microbiol 72: 5254–5259.
Kramarsky-Winter E & Loya Y (1998) Reproductive strategies of
two fungiid corals from the northern Red Sea: environmental
constraints. Mar Ecol Prog Ser 174: 175–182.
Kumar S, Tomura K & Nei M (2004) MEGA3: integrated software
for molecular evolutionary genetics analysis and sequence
alignment. Brief Bioinform 5: 150–163.
Kushmaro A, Loya Y, Fine M & Rosenberg E (1996) Bacterial
infection and coral bleaching. Nature 380: 396.
Lampert Y, Kelman D, Dubinsky Z, Nitzan Y & Hill RT (2006)
Diversity of culturable bacteria in the mucus of the Red Sea
coral Fungia scutaria. FEMS Microbial Ecol 58: 99–108.
Lee YK, Kwon KK, Cho KH, Kim HW, Park JH & Lee HK (2003)
Culture and identification of bacteria from marine biofilms.
J Microbiol 41: 183–188.
Lincoln SP, Fermor TR & Tindall BJ (1999) Janthinobacterium
agaricidamnosum sp. nov., a soft rot pathogen of Agaricus
bisporus. Int J Syst Bacteriol 49: 1577–1589.
Magarvey NA, Keller JM, Bernan V, Dworkin M & Sherman DH
(2004) Isolation and characterization of novel marine-derived
actinomycete taxa rich in bioactive metabolites. Appl Environ
Microbiol 70: 7520–7529.
Maidak B, Cole JR, Parker CT Jr et al. (1999) A new version of the
RDP (Ribosomal Database Project). Nucleic Acids Res 27:
171–173.
Meikle P, Richards NG & Yellowlees D (1988) Structural
investigations on the mucus from six species of coral. Mar Biol
99: 187–193.
Muyzer G, Hottenträger S, Teske A & Wawer C (1996) Denaturing
gradient gel electrophoresis of PCR-amplified 16S rDNA – A
new molecular approach to analyse the genetic diversity of
mixed microbial communities. Molecular Microbial Ecology
Manual (Akkermans ADL, van Elsas JD & Bruijn FJ, eds),
pp. 1–23. Kluwer Academic Publishers, Dordrecht.
FEMS Microbiol Lett 276 (2007) 106–113
113
Coral mucus-associated bacterial communities
Pantos O, Cooney RP, Le Tissier MDA, Barer MR, O’Donnell AG
& Bythell JC (2003) The bacterial ecology of a plague-like
disease affecting the Caribbean coral Montastrea annularis.
Environ Microbiol 5: 370–382.
Paul JH, DeFlaun MF & Jeffrey WH (1986) Elevated levels of
microbial activity in the coral surface microlayer. Mar Ecol
Prog Ser 33: 29–40.
Rappe MS, Vergin K & Giovannoni SJ (2000) Phylogenetic
comparison of a coastal bacterioplankton community with its
counterparts in open ocean and freshwater systems. FEMS
Microbiol Ecol 33: 219–232.
Reshef L, Koren O, Loya Y, Zilber-Rosenberg I & Rosenberg E
(2006) The coral probiotic hypothesis. Environ Microbiol 12:
2068–2073.
Ritchie KB (2006) Regulation of microbial populations by coral
surface mucus and mucus-associated bacteria. Mar Ecol Prog
Ser 322: 1–14.
Ritchie KB & Smith WG (1997) Physiological comparison of
bacterial communities from various species of scleractinian
corals. In: Proc 8th Int Coral Reef Symp Publ 1: 521–526.
Rohwer F, Breitbart M, Jara J, Azam F & Knowlton N (2001)
Diversity of bacteria associated with the Caribbian coral
Montastraea franksi. Coral Reefs 20: 85–91.
Rohwer F, Seguritan V, Azam F & Knowlton N (2002) Diversity
and distribution of coral-associated bacteria. Mar Ecol Prog Ser
243: 1–10.
FEMS Microbiol Lett 276 (2007) 106–113
Rosenberg E, Koren O, Reshef L, Efrony R & Zilber-Rosenberg I
(2007) The role of microorganisms in coral health,
disease and evolution. Nature Rev Microbiol 5:
355–362.
Rosenfeld M, Bresler V &Abelson A (1999) Sediment as a possible
source of food for corals. Ecol Lett 2: 345–348.
Saito N & Nei M (1987) The neighbor-joining method: a new
method for constructing phylogenetic trees. Mol Biol Evol 4:
406–425.
Sambrook J & Russell DW (2001) Molecular Cloning: A
Laboratory Manual, 3rd edn. Cold Spring Harbor Laboratory
Press, Cold Spring Harbor, NY.
Santavy DL & Peters EC (1997) Microbial pests: coral disease
in the western Atlantic. Proc. 8th Int Coral Reef Symp 1:
607–612.
Schloss PD & Handelsman J (2005) Introducing DOTUR, a
computer program for defining operational taxonomic units
and estimating species richness. Appl Environ Microbiol 71:
1501–1506.
Sorokin YI (1973a) Trophical role of bacteria in the ecosystem of
coral reef. Nature (London) 242: 415–417.
Sorokin YI (1973b) On the feeding of some scleractinian corals
with bacteria and dissolved organic matter. Limnol Oceanogr
18: 380–385.
Sorokin YI (1978) Microbial production in the coral-reef
community. Arch Hydrobiol 83: 281–323.
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