Coral mucus-associated bacterial communities from natural and aquarium environments

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

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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., 2007 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved


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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

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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, D¨uren, 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). 2007 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved


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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%

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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, 2007 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved


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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.

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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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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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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 2007 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. All rights reserved


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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.

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