Introduction

Coral reefs are magnificent massive structures covering ∼620 000 km2 of the earth (Farber et al., 2002) and hosting an enormous diversity of organisms (Sebens, 1994). Scleractinian corals, the primary framework builders of coral reefs, comprise a mutualistic interaction between the coral and a dinoflagellate of the genus Symbiodinium, and also contain diverse populations of microorganisms (Bacteria, Archaea, Eucarya and viruses) (Rohwer et al., 2001, 2002; Pantos et al., 2003; Kellogg, 2004; Ritchie and Smith, 2004; Bourne and Munn, 2005; Koren and Rosenberg, 2006) that have apparently co-evolved with the corals (Ritchie and Smith, 2004; Rohwer and Kelly, 2004). This has recently led to the idea that the host (coral animal) and its associated microorganisms should be considered as a holobiont (Rohwer et al., 2002; Rosenberg et al., 2007).

The interaction between corals and their symbiotic microbial community can have a dramatic influence on coral physiology and health. To date, most reports on coral–bacterial interactions have focused on pathogens (reviewed in Harvell et al., 2007). However most bacterial–coral associations are likely to be advantageous, providing essential nutrients for the coral host (for example, nitrogen) (Wegley et al., 2007; Siboni et al., 2008; Rypien et al., 2009; Vega-Thurber et al., 2009) or protecting the coral from infection by producing antimicrobial agents that restrict the growth of potential pathogens (Loya et al., 2001; Ritchie, 2006; Shnit-Orland and Kushmaro, 2009). An interesting hypothesis that was recently presented, and that took into consideration the dynamic relationship between symbiotic microorganisms and corals under different environmental conditions, is the coral probiotic hypothesis (Reshef et al., 2006). This hypothesis suggests that by altering the structure of coral-resident microbial community, the holobiont can adapt to the changing environmental conditions more rapidly and with greater versatility than a process that is dependent on genetic mutation and selection of the coral host (Rosenberg et al., 2007, 2009). This hypothesis is supported by the large diversity in microbial populations associated with corals, the benefit of the coral-associated microbes to the host and the ability to rapidly change the coral-associated microbial population in response to changing environmental conditions.

Industrial revolution, increase in human population and human activities, especially over-fishing and pollution have brought about dramatic and rapid environmental changes that have had substantial influence on the world's oceans. In the last two decades we have experienced an increase in both seawater temperature and carbon dioxide (CO2) concentrations, which are projected to continue to increase over the next 100 years (Hoegh-Guldberg, 1999; Hoegh-Guldberg et al., 2007; Rahmstorf et al., 2007). Climate and environmental changes are having a devastating effect on coral reefs. Reports on coral bleaching and disease outbreaks have been on the rise and several reefs around the world are in danger of extinction (Carpenter et al., 2008). An emerging environmental stress to coral reefs integrity is ocean acidification, a result of increasing atmospheric CO2 concentration. Atmospheric CO2 now exceeds 380 p.p.m., which is more than 80 p.p.m., above the maximum values of the past 740 000 years (Petit et al., 1999). This increase has already resulted in a change in seawater pH (decreased by 0.1 pH unit) since pre-industrial times and is predicted to decrease by up to 0.4 units by the end of the twenty-first century (IPCC, 2007). The ocean absorption of anthropogenic CO2 also reduces the saturation state of seawater with respect to calcite and aragonite, two forms of calcium carbonate secreted by calcifying marine biota. Calcification is strongly dependent on the carbonate saturation state of seawater (Caldeira, 2007). Thus the drastic change in pH may dramatically influence the physiology of corals and other oceanic organisms, in particular organisms that deposit calcium carbonate to build their skeletons/shells (Gattuso et al., 1998; Orr et al., 2005).

Recent studies have characterized the influence of pH decrease on marine ecosystems (Ishimatsu et al., 2005; Bibby et al., 2007; Fine and Tchernov, 2007; Hoegh-Guldberg et al., 2007; Kuffner et al., 2007). These studies presented several effects, such as decreased abundance of crustose coralline algae (Kuffner et al., 2007), shell dissolution in living pteropods (Orr et al., 2005) and chronic effects on fish physiology (Ishimatsu et al., 2005). Fine and Tchernov (2007) reported that a decrease in pH (from 8.2 to 7.4) led to physiology alterations in the coral Oculina patagonica and Madracis pharensis, leading to complete dissolution of their skeleton. It is interesting that the corals returned to calcify when returned to ambient pH after a year in low pH (Fine and Tchernov, 2007).

The change in pH, most likely, does not solely affect the coral host but can also influence the associated microbial community (Vega-Thurber et al., 2009). In this study, we characterized the microbial community associated with the scleractinian coral Acropora eurystoma and the changes in the microbial community following exposure to decreased pH.

Materials and methods

Coral collection and maintenance

The corals used in this study were collected from the reef off the Interuniversity Institute for Marine Sciences in Eilat (IUI, 29°30′N; 34°55′E, Gulf of Aqaba) Red Sea. Twenty colonies of the scleractinian coral Acropora eurystoma were collected at 10–15 m depth and fragmented to 5 cm long fragments. The fragments were immediately placed in running seawater (temperature of 26 °C, salinity 3.9%, irradiance of 250 μmol m−2 s−1 and photoperiod of 10 L:14D and pH 8.2). Following an acclimation period of 2 weeks and full recovery of the tissue, these mini-colonies (obtained from the 20 colonies) were distributed between the two pH treatments: 7.3 and 8.2 and maintained under these conditions for 10 weeks. The distribution of the fragments was carried out systematically such that fragments from the same colony were evenly distributed between the treatments to avoid any possible bias because of sample collection.

To reach the desired pH, water from the Red Sea, pumped from 30 m depth (salinity 39% and temperature 26 °C), was continuously supplied to two 1000 l tanks. pH was manipulated by bubbling pure CO2 into seawater in the tanks and controlled by using a pH controller (Aquastar, IKS ComputerSysteme GmbH, Karlsbad, Germany) connected to pH electrodes (S-200C, Sensorex, Garden Grove, CA, USA). A detailed description of the pH system is provided in the Supplementary Information. All mini-colonies of Acropora eurystoma in the two pH treatments survived to the end of the experiment and did not show any signs of stress.

Coral fractionation, bacteria isolation and DNA extraction

At the end of the experiment (10 weeks) the corals were fractionated to mucus, tissue and skeleton fractions, and samples were taken for isolation of bacteria and DNA extraction. To remove the mucus, the corals were placed in 50-ml centrifuge tubes, centrifuged for 3 min at 2675 g, and the mucus collected. The coral was then washed several times in seawater and the tissue was then extracted from the coral using an airbrush. To minimize any cross contamination from the tissue to the skeleton, the skeleton was thoroughly cleaned from remaining tissue residues by airbrush, followed by several washes with sterile seawater, only then the skeleton was crushed with a mortar and pestle. To isolate bacteria from these fractions, 10-fold serial dilutions of coral mucus, tissue, skeleton and seawater were plated on Marine Agar (Difco, Detroit, MI, USA) and incubated at 26 °C. To obtain bacteria from the seawater, 1 l of seawater from each pH treatment was filtered through polycarbonate filters (0.22 μm). The filters were homogenized in 50 ml of filtered sterile seawater and serial dilutions were plated on Marine Agar. To extract DNA from coral mucus, crushed tissue, skeleton and seawater, samples were centrifuged at 9300 g for 15 min. DNA was extracted from the pellets using the UltraClean Soil DNA kit according to the manufactures guidelines (MoBio, Carlsbad, CA, USA). DNA from cultured bacteria was extracted from individual colonies using a DNA Purification kit (Promega, Madison, WI, USA).

PCR amplification of 16S rRNA gene and clone library construction

Bacterial primers 8F and 1492R (Lane, 1991) were used for amplification of the 16S rRNA gene from the various fractions and isolated colonies. 16S rRNA genes were amplified in a 25 μl reaction mixture consisting of 2.5 μl of 10 × buffer, 2.5 μl of a 2.5 mM total dNTP mixture, each primer at 5 μM, 10 ng of template DNA, and 2.5 U of Ex Taq DNA polymerase (TaKaRa, Bio, Shiga, Japan). Amplification conditions for the PCR included an initial denaturation step of 94 °C for 3 min, followed by 29 cycles of 94 °C for 35 s, 56 °C for 35 s and 72 °C for 1.5 min and a final extension step of 72 °C for 7 min. Reaction products were checked for size and purity on 1% agarose gel. PCR products obtained from bacterial isolates were sequenced. Amplified DNA from seawater, mucus, tissue and skeleton samples was ligated into the pGEM-T Easy vector according to the manufacturer instructions (Promega) and transformed into competent Escherichia coli DH5α. For each fraction, 2–3 libraries were constructed. The clones obtained were amplified by colony PCR with pGEM primers (M13F and M13R). Amplification conditions for the colony PCR included an initial denaturation step of 95 °C for 4.5 min, followed by 29 cycles of 95 °C for 30 s, 59.5 °C for 30 s and 72 °C for 1 min and a final extension step of 72 °C for 10 min. Reaction products were checked for size and purity on 1% agarose gel. Before sequencing, the PCR products were enzymatically purified using ExoSAP-IT (USB, Cleveland, OH, USA). Sequencing was carried in Macrogen, Korea, using the 8F forward primer. The nucleotide sequence data for all of the clones and colonies reported in this paper were deposited in the GenBank nucleotide sequence database under accession numbers GU319121–GU319777.

Phylogenetic analysis

For initial identification the obtained sequences were compared with the NCBI database (National Center for Biotechnology Information; www.ncbi.nlm.nih.gov) using the BLASTn algorithm. Phylogenetic trees were constructed and checked by bootstrap analysis using the ARB phylogenetic program package software (Ludwig et al., 2004). Sequences (650 bp) were aligned to the ARB gene database with the ARB automated alignment tool, and refined manually by visual inspection and secondary-structure analysis. Phylogenetic affiliation of the clone sequences was determined according to both ARB and NCBI-BLAST results. The similarity of the clone libraries from the different fractions was compared by Unifrac Metric analysis using the Jackknife algorithm where 100 replicates of trees were examined (Lozupone et al., 2006). In each replicate tree the different samples were represented by 30 clone sequences each. Diversity indices and operational taxonomic units were determined using DOTUR (Schloss and Handelsman, 2005).

Denaturing gradient gel electrophoresis (DGGE) analysis

PCR amplifications for DGGE were performed as previously described using bacterial primer pair 341F (with a GC clamp at its 5′ end) and 907R (Muyzer and Smalla, 1998). Amplification conditions for the PCR included an initial denaturation step of 95 °C for 4 min, followed by 34 cycles of 94 °C for 30 s, 60 °C for 30 s, 72 °C for 30 s and a final extension step of 72 °C for 10 min. Products were resolved on 6% (w/v) polyacrylamide gels containing a linear urea-formamide gradient ranging from 20 to 60% denaturant (with 100% defined as 7 M urea and 40% formamide). Gels were run for 17 h at 60 °C at a constant voltage of 100 V using the Dcode Universal Mutation System (Bio-Rad Laboratories, Hercules, CA, USA). Community patterns were visualized after staining with Gelstar (Invitrogen, Carlsbad, CA, USA) by UV transillumination (302 nm) and were photographed with a Kodak KDS digital camera (Kodak, New Haven, CT, USA).

Cluster analysis of DGGE community fingerprints

DNA fingerprints obtained from the 16S rRNA gene banding patterns on the DGGE gels were analyzed using FingerprintingII Informatix Software (Bio-Rad Laboratories). The lanes were normalized to contain the same amount of total signal after background subtraction. The gel images were straightened, aligned and analyzed to give a densitometric curve for each gel. Band positions were converted to Rf values between 0 and 1, and profile similarity was calculated by the unweighted pair-group method using arithmetic average algorithm, for the total number of lane patterns. Cluster analyses of the lane patterns were constructed by FingerprintingII software.

Antibacterial screening

Screening the cultured bacteria for antibacterial activity was performed as described previously (Shnit-Orland and Kushmaro, 2009) against common indicator bacteria, including Bacillus subtilis, Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus, as well as the coral pathogen, Vibrio coralliilyticus (Ben-Haim et al., 2003). We used a drop technique for the screening process. Isolated marine bacteria, from the mucus of the corals, were grown on Marine Broth (Difco) for 48 h at 26 °C (ambient seawater temperature). After sufficient growth, a 10 μl drop of the cell suspension (1.0 OD600 nm) was placed on agar plates freshly covered with an overnight culture of a specific indicator bacterium (100 μl of 1.0 OD600 nm culture). Activity was measured after 24 h by the formation of an inhibition zone where the drop had been placed. Each analysis was performed four times. All indicator bacteria were cultured on Luria-Bertani Broth agar (Difco) plates, except for V. coralliilyticus, grown on Marine Broth agar, at their optimal growth temperatures (30 °C for V. coralliilyticus and B. subtilis and 37 °C for the others).

Results

Changes in the microbial community following exposure to decreasing pH

DGGE was initially used to examine the bacterial community composition of the corals exposed to reduced and ambient pH treatments. Pattern comparison (Figure 1) showed that the bacterial communities in all coral fractions (mucus, tissue and skeleton) were influenced by the pH. The clusters were divided according to the pH level that the coral was exposed to (that is, pH 8.2 and pH 7.3) and coral fractions were distributed within the pH clusters, indicating that the bacterial community had shifted in response to the environmental change.

Figure 1
figure 1

Cluster analysis of DGGE patterns of bacterial communities in Acropora eurystoma and seawater exposed to pH 7.3 and 8.2. The cluster analysis was performed using the Fingeprint II software. The unweighted pair-group method using arithmetic average tree was constructed on the basis of Jaccard similarity matrix between the different DGGE patterns and presented in each lane.

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16S rRNA gene clone libraries of A. eurystoma microbial community revealed changes in the community composition following exposure to reduced pH. To further analyze the community composition, 16S rRNA gene clone libraries of A. eurystoma microbial community were constructed and sequenced (n=415). Rarefaction analyses were performed using DOTUR (Schloss and Handelsman, 2005) on the clone libraries to determine the number of unique bacterial clones as a proportion of the estimated total diversity within each library. The rarefaction curves did not reach an asymptote (Supplementary Figure S1), indicating insufficient sampling to capture the total diversity of the bacterial community. Overall we identified 103 operational taxonomic units from the corals maintained at pH 7.3 versus 74 operational taxonomic units in the corals maintained at pH 8.2 (Table 1). To determine the abundance and richness of the bacterial communities associated with the coral, the Shannon–Wiener index was calculated to quantify species diversity (Table 1). The community from pH 7.3 was more diverse than that at pH 8.2, with a Shannon–Wiener index of 4.34 compared with 3.94 at pH 7.3.

Table 1 Number of OTUs and richness estimation of bacterial 16S rRNA gene libraries from corals maintained at pH 8.2 and 7.3
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A phylogenetic comparison between the different treated samples showed considerable differences in the microbial community composition (Figure 2a) that closely reflected the clustering observed with DGGE analysis. The clusters were mainly divided by the pH treatments and not by the coral fractions (that is, mucus, tissue and skeleton). Furthermore, there was a clear difference between the bacterial community in the coral and that of the surrounding water.

Figure 2
figure 2

Cluster analysis of sequences obtained from microbial community 16S rRNA clone libraries from A. eurystoma (a) uncultured 16S rRNA clones (n=415). (b) Cultured isolates (n=263). *Note that the isolates from skeletons of corals maintained at pH 7.3 are clustered with isolates from the corals maintained at pH 8.2 (see text). Cluster analysis was performed using Jackknife Environment Clusters with 100 permutations and minimum sampling size of 30 sequences per environment (Unifrac Metric analysis) (Lozupone and Knight, 2005).

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Diversity of bacterial groups and their distribution

Sequences obtained from the 16S rRNA gene libraries were used to analyze the diversity of bacterial communities associated with A. eurystoma exposed to pH 8.2 and 7.3 (Figure 3 and Supplementary Table S1 showing the percent distribution between fraction in each treatment, Supplementary Table S2 providing the complete list of clones analyzed in the study and Supplementary Tables S3–S4 providing the most abundant clones in each treatment). The two dominant bacterial groups Gammaproteobacteria (26 and 27% of the total clones at pH 8.2 and 7.3, respectively) and Cyanobacteria (23 and 38%, respectively) remained dominant in the corals incubated at the two pH treatments (Figures 3a and b). Furthermore in several instances a close homolog of the same strain was present at both pH treatments, for example: Leptolyngbya sp. (EU249128), Phormidium persicinum (EF654085) and Thalassolituus sp. (GQ426888) (Supplementary Tables S2–S4). A significant change in gene frequency was observed in the following bacterial groups at the lower pH. The Alphaproteobacteria increased in their overall presence from 11 to 20% and the dominant clones belonged to the Rhodobacteraceae. Two other groups in which their gene frequency had changed were Deltaproteobacteria and Bacteroidetes. The gene frequency of Deltaproteobacteria was reduced from 24 to 5% when shifted to pH 7.3. This group was especially dominant in the skeleton fraction (see Supplementary Tables S1–S4) and most bacteria in this group (more than 60%) showed homology to Desulfobacter species that belong to the sulfate-reducing bacteria. The gene frequency of the Bacteroidetes decreased from 6 to 2%. Examination of the community structure within the fractions (mucus, tissue and skeleton) further reveals these trends (Supplementary Tables S1 and S2).

Figure 3
figure 3

The diversity of the bacterial groups found in A. eurystoma and seawater exposed to pH 7.3 and pH 8.2. The chart was constructed on the basis of 16S rRNA gene sequences derived from clone libraries obtained from the coral A. eurystoma maintained at pH 7.3 (n=173) or 8.2 (n=159) and from the surrounding water “sw” maintained at pH 7.3 (n=43) and 8.2 (n=24). The alpha, beta, gamma, delta and epsilon belong to the Proteobacteria phylum.

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Changes were also detected in the less abundant bacterial groups. Of those, noteworthy are the changes in Verruncomicrobiae and Gammatimondetes, which were present only at pH 7.3, whereas other groups, such as Epsilon and Beta Proteobacteria, were only present in the corals maintained at ambient pH (Figure 3 and Supplementary Tables S1 and S2). The coral microbial community showed a clear difference from the microbial community of the surrounding water (Figures 2a and 3 and Supplementary Table S2). Furthermore, no major difference in diversity was evident between the surrounding water maintained at the two pH levels (Figures 2a and 3c, d).

Diversity of the culturable bacterial communities and their antibacterial activities

The diversity of bacteria isolated from the coral fractions exposed to the two pH conditions was examined. Analysis of the 16S rRNA gene sequences obtained from isolates (n=263) showed that the clustering into two different communities correlated, to some extent, with pH treatment (Figure 2b). One cluster contained isolates obtained from mucus and tissue fractions from the coral maintained at pH 7.3. The other contained the bacteria isolated from the mucus, tissue and skeleton at pH 8.2. It is interesting that the skeleton bacterial community isolated from corals at pH 7.3 clustered with the bacterial communities fractions isolated from corals exposed to pH 8.2 (Figure 2b). As expected, the diversities of the culturable isolated bacteria from both pH treatments differed from those of the culture-independent groups (Figures 3 and 4 and Supplementary Tables S1–S4). For example, Bacillus sp. (Bacillus jeotgali (GU397390) and Bacillus litoralis (FJ188307)) were dominant in the culturable fraction and rarely detected in the culture-independent group. Similarly, Rohodobacteraceae were dominant at pH 8.2 in the culturable fraction and decreased in pH 7.3. This was opposite to the trend that was evident for this family in the culture-independent group.

Figure 4
figure 4

The diversity of the bacterial groups isolated from A. eurystoma exposed to pH 7.3 and pH 8.2. The chart was constructed on the basis of 16S rRNA gene sequences derived from bacterial isolates that were associated with A. eurystoma maintained at pH 7.3 (n=150) and 8.2 (n=108).

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An important difference apparent in the isolated community at the lower pH treatment was the increase in Vibrionaceae (for example, Vibrio sp. GQ406613 and uncultured Vibrio sp. AM941184) and Alteromonadaceae (Alteromonas sp. GU726843) (Tables S2–S4). In a few cases the isolates were detected in corals maintained at pH 8.2, but increased in pH 7.3, for example Vibrio sp. (FJ161306) and Alteromonas sp. (GU726843) (see Supplementary Table S2). This change is especially interesting as species that belong to Vibrionaceae and Alteromonadaceae were previously associated with diseased or stressed corals (Ritchie, 2006; Bourne et al., 2007; Arboleda and Reichardt, 2009; Sunagawa et al., 2009).

To examine the antibacterial activity of isolated bacteria from each pH, a recently established screening method was used (Shnit-Orland and Kushmaro, 2009). The antibacterial activity of 165 isolates was tested against five indicator strains (Bacillus subtilis, Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus, as well as the coral pathogen Vibrio coralliilyticus). The screen was carried out with bacteria isolated from the mucus fraction, as we expected these bacteria to most likely serve as the first line of defense. Corals maintained at pH 7.3 showed a marked increase in the isolated bacteria, producing antibacterial activity (42%) as compared with isolates obtained from corals maintained at the ambient pH (17%). It is interesting that most of the bacteria that showed antibacterial activity belong to the Vibrionaceae and Rhodobacteraceae, 50 and 29%, respectively. The complete list of bacteria showing antimicrobial activity and against which indicator strains are summarized in Supplementary Table S5.

Discussion

Coral colonies represent a complex landscape, comprising the coral animal, a community of symbiotic dinoflagellates and associated bacterial biota that can thrive in fascinating harmony. Many environmental factors can influence this delicate balance and affect coral health and disease (Bruno et al., 2007; Rosenberg et al., 2007; Garren et al., 2009). This complexity also provides means for coral resistance and persistence. One of the major environmental factors that may directly affect coral health and this delicate symbiosis is an increase in CO2 concentration, which is projected to result in ocean acidification (Anthony et al., 2008). Extensive research is currently on-going to examine the impact of changes in pH on the physiology of corals, but little is known about the impact of reduced pH on coral-associated microbial communities (Vega-Thurber et al., 2009). The current study addresses this question by examining whether or not reduced pH can influence the microbial community structure and ability to provide ecological services to the coral host (for example, antibacterial activity). It should be emphasized that the pH used in this study is lower than what is expected in the next 100 years.

The results show that the coral A. eurystoma harbors a specific microbial community that is distinct from the surrounding water, as reported in other corals from various locations (Bourne and Munn, 2005; Guppy and Bythell, 2006; Ritchie, 2006; Kooperman et al., 2007; Lampert et al., 2008; Arboleda and Reichardt, 2009). The coral–bacterial community structure was influenced by seawater pH, and the communities were clustered according to the pH treatment, more so than the coral micro-environment, for example, mucus, tissue and skeleton. A higher Shannon–Wiener index, indicating an increase in coral-associated microbial diversity, was recorded at the lower pH treatment. This could be the result of an intermediate disturbance (Connell, 1978), leading to an increase in biodiversity. We speculate that reduced pH triggers this disturbance. When characterizing the microbial community of the A. eurystoma using 16S rRNA clone libraries, the major bacterial groups found were similar to those found in other studies (Sekar et al., 2006; Garren et al., 2009; Nithyanand and Pandian, 2009; Rypien et al., 2009; Vega-Thurber et al., 2009). These included mainly the Gammaproteobacteria and Cyanobacteria, which composed approximately 50% of the bacterial community and remained dominant in the corals incubated at the two pH treatments. However, several interesting shifts in the microbial community were found following the decrease in pH. The Alphaproteobacteria, dominated by Rhodobacteraceae, increased in their overall presence from 11 to 20% (pH 8.2 and 7.3, respectively). Sequences found from this group were closely related to sequences that were previously described in studies examining the microbial community associated with diseased corals, such as white plague disease in Montastraea faveolata (GenBank accession numbers FJ403086, FJ202287, FJ202842, FJ203583, FJ202745 and FJ203565 all showing 97–98% sequence identity), white syndrome in Turbinaria mesenterina (GenBank accession number EU780231, 85–91% identity) and Muricea elongate in bleached corals (GenBank accession number DQ917824, 98% identity). In addition, clones reported to be associated with Oculina patagonica (Koren and Rosenberg, 2006) and sponge-associated bacteria isolated from Axinella corrugate (GenBank accession number EF092256, 99% identity) and Haliclona (gellius) sp. (GenBank accession number U346416, 92% identity) were also detected.

The Deltaproteobacteria, which contained sequences primarily homologous to species belonging to the sulfate-reducing bacteria Desulfobacter, had a lower frequency (5%) at pH 7.3 as compared with pH 8.2 (24%). sulfate-reducing bacteria have been implicated as part of the microbial consortium that induces black band disease in corals (Schnell et al., 1996; Richardson et al., 1997) or to increase in frequency in corals exposed to pollution (Garren et al., 2009). No signs of disease were detected in the corals (at either pH), and it is important to note that sulfate-reducing bacteria are also present in healthy corals (Arboleda and Reichardt, 2009).

The abundance of the Bacteroidetes decreased from 6% (pH 8.2) to 2% (pH 7.3). In this group, clones were found related to coral bacterial symbionts previously found in the Caribbean coral Montastraea faveolata (Sunagawa et al., 2009), Siderastrea siderea (Sekar et al., 2008) and the Mediterranean coral Oculina patagonica (Koren and Rosenberg, 2006). Although Bacteroidetes have been previously detected in healthy corals (Sekar et al., 2008), an increase in Bacteroidetes was associated with Black Band Disease (Cooney et al., 2002; Frias-Lopez et al., 2002, 2004; Barneah et al., 2007). Bacteroidetes were also reported to increase in corals exposed to various stress conditions (Vega-Thurber et al., 2009).

Another interesting finding we observed when examining the community in the corals exposed to pH 7.3 was the increase in species showing close homology to families that include known coral pathogens, such as Vibrionaceae and Alteromonadaceae (Ritchie, 2006; Bourne et al., 2007; Arboleda and Reichardt, 2009; Sunagawa et al., 2009), as well as Rhodobacteraceae (Alphaproteobacteria), previously isolated from diseased, injured or stressed marine invertebrates (Sekar et al., 2006; Sunagawa et al., 2009).

Anthony et al. reported an adverse effect of reduced pH on corals health and function (Anthony et al., 2008). In the present study, we did not observe signs of bleaching or disease and did not detect identical sequences of known coral pathogens. Close homologous of bacteria reported to be associated with diseased or stressed corals were detected in the lower pH treatment. For example, bacteria isolated from a diseased gorgonian coral showing homology to the coral pathogen V. coralliilyticus were identified (GQ406701, 93% identity and GQ406613, 98% identity). In addition, close homologous of bacteria associated with diseased marine organisms were also identified (for example, Vibrio pectenicida a pathogen of scallop (Y13830, 98% identity) and bacteria isolated from diseased fish (FN432989, 99% identity)). Part of this change in the bacterial community was markedly detected only in the culturable bacteria (Vibrionaceae and Alteromonadaceae), yet we believe this trend represents an important phase-shift in the community, probably a result of environmental change and/or changes in the physiological state of the holobiont. Several studies have reported an increase in similar bacterial families under various stress conditions. For example, Ritchie examined mucus from healthy Acropora palmata during high temperatures and reported that Vibrio sp. increased and replaced the community of beneficial bacteria (Ritchie, 2006). Garren et al. (2009) also observed an increase in the number of Vibrios following exposure of corals to yet another common stressor increased pollution as a result of intensive fish farming (Garren et al., 2009). Similar results were also reported in a recent study examining the metagenomic profile of coral holobionts exposed to various stressors (Vega-Thurber et al., 2009). It is interesting that one of the stress conditions tested in that study was pH. Although the coral species was different (Porites compressa), the pH was lower (6.7 versus 7.3) and the exposure time shorter (64 h versus 10 weeks in our study), the authors also reported that decreased pH resulted in a shift from a bacterial community associated with healthy corals towards a community often associated with diseased corals. Vibrio sp., for example, increased at the lower pH treatment and were found to significantly alter the microbiome metabolism (Vega-Thurber et al., 2009). On the basis of this, the authors hypothesized that the contribution of just a few members of a community can profoundly shift the health status of the coral holobiont (Vega-Thurber et al., 2009). It is hard to conclude from our study whether the corals indeed experienced environmental stress due to the decrease in pH. However, the shift to opportunistic bacterial species that are known to be associated with diseased or stressed corals is in our opinion an important biomarker that may suggest this is indeed the case. Our study opens the path for future work that can address this important hypothesis.

Changes in bacterial communities usually provide limited information on the direct impact the microbial community may have on the coral host. One of the important traits that is thought to be contributed by the microbial community and can influence the coral host is the production of antibacterial agents (Kelman et al., 2006; Nithyanand and Pandian, 2009; Rypien et al., 2009; Shnit-Orland and Kushmaro, 2009). We examined the ability of the isolated bacterial community to produce antibacterial activity against five bacterial species (including a coral bleaching pathogen) and detected an increase from 17 to 42% in the percent of coral bacteria showing antibacterial activity following a decrease in pH. This increase in antibacterial activity may impact the dynamics of the microbial community and highlights the forces that take place in structuring its diversity. In fact, most of the antibacterial activity was produced by isolates belonging to the Vibrio sp. (50%), suggesting that this may be one of the mechanisms by which these bacteria may colonize the coral host. These results further support the metagenomic study by Vega-Thurber et al. (2009) who reported that under low pH, in addition to the increase in pathogens, there was an increase in secondary metabolites and stress resistance (for example, antibiotics and toxins) (Vega-Thurber et al., 2009).

In conclusion, the present study emphasizes the complexity of the coral holobiont and its response to changes in environmental conditions. The microbial community may be a key factor in determining holobiont's health and function. It is, therefore, imperative to further explore interactions between the microbial community and its hosting coral under environmental change, such as pH.