Abstract
Gorgonian corals are among the most representative species of hard-bottom sublittoral ecosystems in the Mediterranean Sea. The red gorgonian, Paramuricea clavata, harbors a specific, spatially and temporally stable bacterial community that may have an important biological role in the function of the holobiont. However, little is known about the microbial–gorgonian relationship, specifically the nature of the interaction established by the bacteria with their host. Documenting the distribution of microorganisms within coral tissue compartments is critical for providing an overall framework for understanding their functional roles. Here, we used histology and fluorescence in situ hybridization (FISH) to detect and localize bacteria in the tissues of P. clavata. Aggregates of tightly packed bacteria were found in the endoderm of the polyps, which is consistent with previous observations of endobacteria in the tissues of scleractinian coral species. This finding suggests the existence of an intimate biological integration between specific members of the microbiota and gorgonian corals that likely results from a complex interplay mediating recognition and maintenance of the bacterial associates. In addition to providing the basis for the development of FISH studies in gorgonians, this work includes the first histological description of the inner structure of P. clavata polyps, illustrating the typical features of general gorgonian anatomy.
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Introduction
Corals are associated with diverse assemblages of bacteria that are believed to contribute to their overall health and adaptive response to environmental changes (Rosenberg et al. 2007; Bourne et al. 2009). Among other putative functions, coral-associated bacterial communities likely play substantial roles in supporting the host’s nutrient budget and defense against pathogens (for reviews, see Mouchka et al. 2010; Krediet et al. 2013). Despite growing evidence of a critical contribution of bacterial interactions to the healthy functioning of the coral holobiont, relatively little is known regarding the nature of these associations and the spatial distribution of the microbiota within the coral host. Therefore, investigating the architecture of bacterial communities is essential to characterize the microbial processes performed by the natural assemblages in situ and to understand the mechanisms underlying their maintenance within the coral niche. Given increasing stress factors and the worldwide decline of coral health (Hughes et al. 2010), such information could also yield valuable insights into the conditions that may cause the destabilization and breakdown of the holobiont.
In the Mediterranean Sea, gorgonian corals (Cnidarian, Octocorallia) are important structural organisms in coralligenous outcrops, which are one of the most diverse coastal ecosystems (Ballesteros 2006). Among the emblematic long-lived species typically found in coralligenous assemblages is the red gorgonian, Paramuricea clavata, a temperate, azooxanthellate octocoral that forms dense populations in dim-light habitats. Culture-independent surveys of the composition and dynamics of the bacterial communities associated with P. clavata have revealed the presence of host-specific bacterial associates that are conserved throughout seasons and among different geographic locations. These surveys provided a first insight into the microbiology of this species (La Rivière et al. 2013). Recent phylogenetic analyses have also indicated that the dominant bacterial taxa associated with P. clavata and several other gorgonian species may coevolve with their host, suggesting an intimate partnership within the gorgonian holobiont (La Rivière et al. 2015). In addition to genetic-based analyses, further research, such as the study of the spatial arrangement of bacteria within gorgonian tissues, is needed to delineate their potential role and the basis of host specificity.
Fluorescence in situ hybridization (FISH) allows direct observation and identification of bacterial cells in complex natural samples without prior cultivation (Amann et al. 1990b; Amann and Fuchs 2008). This technique has been successfully applied to visualize bacteria associated with different marine invertebrates, such as sponges (Webster et al. 2001), bivalves (Jensen et al. 2010) or crustaceans (Meziti et al. 2010). However, relatively few studies have investigated the localization of individual bacterial cells in corals, notably because coral tissues exhibit a strong endogenous fluorescence that hinders their detection by FISH (Salih et al. 2000; Dove et al. 2001). In certain instances, this problem has been circumvented through improvement in the FISH probe–fluorochrome detection step (e.g., Ainsworth et al. 2006). In particular, Neulinger et al. (2009) applied a catalyzed reporter deposition (CARD)–FISH technique that provides a substantial increase in fluorescence intensity and allows more sensitive detection of the hybridized bacteria in coral tissues (Bobrow et al. 1989; Pernthaler et al. 2002). Therefore, the improvement in direct imaging techniques in future studies will reveal increasing details of the spatial arrangement of coral-associated bacteria.
In this study, our specific objectives were to detect and localize the bacterial associates in the tissues of P. clavata through the adaptation of a CARD–FISH protocol. Because there is currently limited information regarding the internal anatomy of this species, we also aimed to describe the gross organization of the polyp tissues using a histological analysis of stained sections. To the best of our knowledge, this is the first report to provide structural observations of this gorgonian species. The present study will help lay the foundation for future detailed analyses of the nature and significance of the microbial interactions in a temperate gorgonian holobiont.
Materials and methods
Sampling, fixation and decalcification
Apical fragments (2 cm in length) of P. clavata were sampled from three colonies at the Riou site (France; 43°10.345′N, 05°23.319′E) in May 2012 as previously described (La Rivière et al. 2013). The samples were soaked in three washes of sterile-filtered seawater to remove loosely attached bacteria and left in a final bath for 4 h at 15 °C to allow polyp extension. Menthol crystals were added to anesthetize the polyps during a further incubation of 2 h at 4 °C (Lonkar et al. 2011) prior to fixation of the samples in 4 % formalin for 16 h and decalcification in 20 % (wt/vol) EDTA with PBS (pH 7.4) for 48 h at 4 °C. Fragments were subsequently transferred to a 50 % (vol/vol) ethanol–PBS solution (pH 7.4) and stored at −20 °C until use.
Thin sectioning
The fixed tissues were sequentially processed through 70, 80 and 95 % ethanol followed by three 100 % ethanol and three xylene washes, for 1 h each, prior to paraffin embedding. Tissue sections (5–10 µm) were deparaffinized by washing in two baths of Neo-Clear (Merck) for 15 min each and rehydrated in a graded ethanol–water series (twice in 100 % ethanol, once in 95, 90 and 80 % [vol/vol]); they were washed twice in sterile ultra-pure water for 5 min each and then air-dried.
For the light microscopy observations, the sections were stained with a standard hematoxylin and eosin protocol (Sigma-Aldrich) or a Gram staining kit (Sigma-Aldrich) according to the manufacturer’s instructions. For fluorescence in situ hybridization, the slides were processed as described below.
Permeabilization
Bacterial cell walls were permeabilized by incubating the slides in 80 µl of lysozyme solution (1355.106 U of lysozyme [Sigma-Aldrich] ml−1, 500 mM EDTA [pH 8.0], 300 mM Tris–HCl [pH 8.0]) at 37 °C for 2 h in a humidified chamber. After three 1-min washes with sterile ultra-pure H20 at room temperature, the samples were permeabilized with 80 µl of achromopeptidase buffer (60 U of achromopeptidase [Sigma-Aldrich] ml−1, 10 mM NaCl, 10 mM Tris–HCl [pH 8.0]) at 37 °C for 1 h. Subsequently, the slides were washed with sterile ultra-pure H20 and air-dried. Endogenous peroxidases were inactivated by incubating the thin sections in 80 µl of 10 mM HCl solution for 30 min, followed by three washes with sterile H20.
Autofluorescence reduction
The permeabilized slides were buffered in PBS (pH 7.4) for 3 min, followed by 3 min in 70 % ethanol. The thin sections were then covered with Autofluorescence Eliminator Reagent (Chemicon–Millipore) and placed in a humidified chamber overnight (14–16 h). After incubation, the slides were washed three times in 70 % ethanol to ensure complete elimination of the dark reagent, air-dried and stored at −20 °C before hybridization.
Hybridization
Four horseradish peroxidase-labeled oligonucleotide probes were used: the universal eubacterial probes EUB338, EUB338-II and EUB338-III in an equimolar mix (Amann et al. 1990a; Daims et al. 1999) and the control probe non-EUB338 complementary to EUB338 (Wallner et al. 1993). The thin sections were covered with a hybridization solution containing 1 ng µl−1 of probe in hybridization buffer. The hybridization buffer consisted of 55 % formamide, 900 mM NaCl, 20 mM Tris–HCl (pH 8.0), 0.05 % Triton X100 (Sigma-Aldrich), 10 % (wt/vol) dextran sulfate (Sigma-Aldrich) and 10 % blocking solution. The blocking solution (pH 7.5) consisted of 10 % (wt/vol) Blocking Reagent (Boehringer Mannheim), 100 mM maleic acid (Sigma-Aldrich) and 150 mM NaCl. The hybridization was performed at 35 °C for 4 h in a humid chamber. Thereafter, the sections were transferred to 50 ml of prewarmed washing buffer (13 mM NaCl, 20 mM Tris–HCl [pH 8.0], 5 mM EDTA [pH 8.0] and 0.01 % [wt/vol] sodium dodecyl sulfate) at 37 °C for 15 min. The slides were soaked for a few seconds in cold sterile ultra-pure water and quickly air-dried.
Amplification
The thin sections were equilibrated with PBS (pH 7.4) for 15 min in the dark. The tissues were covered with a 1:100 (vol/vol) mixture of tyramide-Alexa488 and amplification buffer (10 % [wt/vol] dextran sulfate [Sigma-Aldrich], 2 M NaCl, 10 % blocking solution, 0.0015 % H202 and PBS [pH 7.4]) and incubated at 46 °C for 30 min in a dark humidified chamber. After amplification, the slides were washed in PBS (pH 7.4) for 20 min at room temperature and three times for 1 min each in sterile ultra-pure water. The sections were air-dried and covered with a DAPI mix mounting medium that allowed counterstaining of the cells (1 µg ml−1 DAPI [4′,6′-diamidino-2-phenylindole], 77 % [vol/vol] Citifluor AF1 [Citifluor Ltd], 14 % [vol/vol] Vectashield mounting medium [Vector Laboratories]).
Microscopy and image processing
The hybridized thin sections were viewed using a Zeiss LSM 780 confocal microscope. The fluorescence was excited sequentially with the 405 and 488 nm laser lines, and the emission was detected in the range from 415 to 469 nm for DAPI and from 497 to 552 nm for Alexa488. Image stacks (steps of 0.5–1 µm) were acquired with GaAsP photomultiplier tube (PMT) detectors. The 32 channels GaAsP PMT detectors allowed spectral detection in parallel mode in the range from 412 to 692 nm. Composite and overlay images were produced from these stacks with ImageJ, v1.46 software (http://imagej.nih.gov/ij).
Results and discussion
Because the histological descriptions of gorgonian corals are scarce, a preliminary analysis of the P. clavata anatomy was needed as a prerequisite for the in situ localization of bacteria. Microscopic examination of hematoxylin–eosin-stained sections demonstrated that the inner structure of P. clavata was in agreement with the previously described anatomy of gorgonians (Galloway et al. 2007). All the examined polyps from the sampled colonies were retracted or moderately expanded, indicating an incomplete effect of the anesthesia procedure. A longitudinal section of polyps retracted in their calyx is shown in Fig. 1a. The central axis consisted of a wide, apparently hollow central core surrounded by concentric layers of proteinaceous gorgonin, which is consistent with the morphology described for other members of the Plexauridae family (Bayer 1981; Lewis et al. 1992). Gonads of different sizes (approximately 100-300 µm in diameter) were found in the polyp cavity of the analyzed sections. In the transverse sections, the pinnate tentacles surrounding the mouth were clearly observed (Fig. 1b). Within the mesoglea, a large number of openings were visible corresponding to the lacunae that remained after decalcification of the sclerites embedded in the host tissue (Nonaka et al. 2012). The mesenteries dividing the gastric cavity could also be discerned (Fig. 1b). Therefore, we conclude that the structural integrity of P. clavata tissues, which was required for in situ localization of bacterial associates, was maintained throughout the steps of our histological protocol.
In a first attempt to determine the presence of bacteria within the tissues of P. clavata, we examined Gram-stained sections of polyps using light microscopy. Putative bacterial aggregates with a round to ovoid shape were consistently observed within the gastrodermal tissue lining the gastric cavity (Fig. 1c, d). The aggregates stained Gram-negative, and their sizes ranged from approximately 10 to 25 µm in maximum diameter. Similar bacterial aggregates were observed in the tissue of several species of scleractinian corals using histology and light microscopy (Sudek et al. 2012; Work and Aeby 2014). Notably, microbial aggregates were also found in the tissues of sea anemones (McKinstry et al. 1989; Schuett et al. 2007), indicating that such intimate bacterial–host associations may be a general feature of cnidarians. In the P. clavata sections, bacteria-like forms could be discerned within certain aggregates and appeared as densely packed spheres of approximately 1–2 µm in diameter. Additional research using electron microscopy is necessary to describe the morphology of the bacteria and their arrangement within the aggregates. Using the Gram stain, no bacteria were detected in other portions of the polyps, such as in the gonads or in the peripheral surface tissues of the gorgonian corals.
Previous attempts to visualize the coral-associated bacterial communities using the FISH procedure have been limited by the endogenous fluorescence of coral tissues (Salih et al. 2000; Ainsworth et al. 2006). A preliminary analysis of P. clavata sections also revealed a high level of autofluorescence that precluded detection of the specific labeling probe (not shown). To overcome this limitation, we resorted to a tyramide-based amplification reaction (CARD-FISH) designed to increase the signal intensities of hybridized bacteria (Bobrow et al. 1989; Pernthaler et al. 2002; Neulinger et al. 2009) and improved the protocol by incubating the tissue sections with an autofluorescence eliminator reagent before the in situ hybridization. This method resulted in a substantial decrease in the background autofluorescence of gorgonian tissues for a range of wavelengths that corresponded to the emission of the FISH probe fluorochromes (Fig. 2).
The P. clavata tissue sections were hybridized with a combination of EUB338 probes that target the majority of the Bacteria domain (Daims et al. 1999). Bright fluorescence signals were observed in the endodermal tissues surrounding the gastric cavity and within the proximal regions of the tentacles (Fig. 3a). The fluorescent bacteria were packed in ovoid aggregates of approximately 12-20 µm in length. Based on their size and shape, these aggregates resembled the Gram-stained bodies within the gastrodermal tissues of P. clavata observed under light microscopy (Fig. 1c, d). However, our imaging technique was not sensitive enough to visualize single fluorescent bacterial cells, precluding the observation of their arrangement within the aggregates. The hybridized sections of P. clavata were also counterstained with DAPI to confirm that the structures detected upon hybridization with the EUB338 probes corresponded to DNA-containing cells (Fig. 3b). No other bacterial signals could be observed on the peripheral, outer surface of the cross-sectioned polyps, thus indicating that the gorgonian microbial community was primarily composed of endobacteria.
Previous studies using FISH documented bacterial aggregates associated with the gastrodermal tissue of several species of scleractinian corals (Ainsworth et al. 2006; Ainsworth and Hoegh-Guldberg 2009). In two coral species from the Great Barrier Reef, Acropora aspera and Stylophora pistillata, the bacterial aggregates within the gastrodermis consisted of Gammaproteobacteria (Ainsworth and Hoegh-Guldberg 2009). Further FISH analysis revealed that tissue-associated aggregates in S. pistillata are composed of members of the Endozoicomonas genus, within the Oceanospirillales order (Bayer et al. 2013b). Recently, Endozoicomonas-related ribosomal sequences have been isolated from an increasing number of healthy coral species (Hansson et al. 2009; Kvennefors et al. 2010; Morrow et al. 2012; Speck and Donachie 2012). Significantly, we have shown that the bacterial community associated with P. clavata is largely dominated by gammaproteobacterial sequences that belong to Endozoicomonas-related bacteria (La Rivière et al. 2013). Endozoicomonas have also been identified as dominant bacterial associates in two other Mediterranean gorgonians, Eunicella cavolini and Eunicella singularis (Bayer et al. 2013a; La Rivière et al. 2015). Thus, the fluorescent aggregates identified by CARD–FISH in P. clavata tissues most certainly correspond to the prevalent Endozoicomonas bacterial associates. However, additional FISH studies with group-specific probes are needed to assess their taxonomic identification and examine their presence within the tissues of other gorgonian species.
Despite accumulating evidence that natural coral-associated populations such as Endozoicomonas spp. contribute to the maintenance of overall coral health (Meyer et al. 2014; Lee et al. 2015; Tout et al. 2015b), their precise functional role is not known. Bacterium–animal endosymbiosis is often related to nutritional interactions with the eukaryotic partner and may be essential to complement the metabolic networks of the host (Moya et al. 2008). Bacteria belonging to the Oceanospirillales order enable heterotrophic degradation of various organic substrates by producing extracellular hydrolytic enzymes (Garrity et al. 2005). Notably, bone-eating marine worms of the genus Osedax harbor intracellular Oceanospirillales that might help their host exploit complex carbon sources such as collagen from whale bones (Goffredi et al. 2007). It is thus tempting to speculate that the localization of bacteria in the coral gastrodermis could be related to nutrient acquisition. Tropical corals and temperate gorgonians are suspension feeders that capture and ingest zooplanktonic prey such as copepods and nauplii (Coma et al. 1994). The production of degradative enzymes by the resident population of endobacteria could therefore aid the digestion of recalcitrant compounds originating from zooplankton and contribute to supply the host with essential nutrients that have been reported to accumulate in the coral gastric fluid (Agostini et al. 2012). Nutrient availability is probably an important constraining factor for the ecology of azooxanthellate gorgonians, such as P. clavata, that cannot beneficiate metabolites from algal partners and are strongly dependent on feeding to sustain their metabolism (Coma et al. 1994). Because food availability is also a critical determinant of the survival of P. clavata during thermal stress (Coma et al. 2009), the possible trophic interaction between the endobacteria and their host may contribute to the physiological adaptation of gorgonians and their resilience to adverse environmental conditions. Potential beneficial roles of Endozoicomonas-related bacteria other than food digestion have been suggested in sulfur recycling (Raina et al. 2009), carbohydrate metabolism (Ding et al. 2016) and production of antibacterial compounds (Rua et al. 2014).
Based on histological slides, all sampled colonies produced female gonads, and no sperm sacs (male gonads; Coma et al. 1995) were observed. No fluorescent bacteria were detected in the oocytes of the examined polyps (Fig. 3c, d), which suggests that P. clavata does not transmit bacteria vertically via gametogenesis and spawning. Although further research (e.g., hybridization of male colonies) is required, this observation is consistent with previous studies showing that spawning corals acquire their bacterial associates via horizontal uptake during early developmental stages, after the release and fertilization of larvae (Apprill et al. 2009; Sharp et al. 2010). Assuming that P. clavata bacterial associates are acquired each generation from the surrounding environment, their establishment within tissues likely depends on complex processes involving selective attraction, recognition and entry steps (Bright and Bulgheresi 2010). Chemoattraction to various compounds released by the corals can induce free-living bacteria to move toward their host, resulting in physical contact between the two partners. Chemotaxis and motility functions identified in coral-associated bacteria, including Endozoicomonas spp., might contribute to this behavior (Tout et al. 2015a). Some mechanism of recognition must exist to determine which bacteria can enter coral tissues. In scleractinian corals, the establishment of algal symbionts within the gastrodermal tissues has been suggested to involve components of the coral immune innate repertoire, such as lectins that may play a role in the recognition and maintenance of Symbiodinium spp. (Wood-Charlson et al. 2006). Because coral lectins also exhibit the ability to bind a variety of bacteria (Kvennefors et al. 2008), we can speculate that the cellular process leading to the entry of bacterial endosymbionts within P. clavata tissues may be closely related to the uptake of Symbiodinium and rely on the octocoral innate immune system (Burge et al. 2013). Additional host-driven factors, such as the production of selective antimicrobial compounds, might contribute to recruit stable and specific symbionts from the environment (Franzenburg et al. 2013). Bacterial endosymbionts might have also evolved various strategies to colonize their particular host partner (Bright and Bulgheresi 2010). Consistent with the possibility of an intimate coadaptation between coral host and symbiont (La Rivière et al. 2015), the recent genomic analysis of a coral-associated Endozoicomonas bacterium suggested its ability to interfere with host cellular signaling to facilitate the entry into tissue layers (Ding et al. 2016).
In conclusion, this study documents the spatial distribution of tissue-associated bacteria in the gastrodermis of a gorgonian coral and may provide a valuable guide to further address the nature of interactions between endobacteria and their host. The consistent retrieval of similar bacterial associations within the gastrodermal tissues of various coral species suggests that endobacteria are an important component of the holobiont and may supply their host with additional nutritional capabilities. Future exploration of the coral–endobacterial relationships would benefit from a combination of in situ visualization approaches, such as CARD–FISH and isotopic tracing (Pett-Ridge and Weber 2012), to investigate host–microbe nutrient exchanges and their potential contribution to holobiont physiology. Such information may be essential to understand the factors mediating the establishment of endosymbiotic bacteria in coral tissues and how they influence the coral fitness in response to environmental changes.
References
Agostini S, Suzuki Y, Higuchi T, Casareto BE, Yoshinaga K, Nakano Y, Fujimura H (2012) Biological and chemical characteristics of the coral gastric cavity. Coral Reefs 31:147–156. doi:10.1007/s00338-011-0831-6
Ainsworth TD, Hoegh-Guldberg O (2009) Bacterial communities closely associated with coral tissues vary under experimental and natural reef conditions and thermal stress. Aquat Biol 4:289–296. doi:10.3354/ab00102
Ainsworth TD, Fine M, Blackall LL, Hoegh-Guldberg O (2006) Fluorescence in situ hybridization and spectral imaging of coral-associated bacterial communities. Appl Environ Microbiol 72:3016–3020. doi:10.1128/AEM.72.4.3016-3020.2006
Amann R, Fuchs BM (2008) Single-cell identification in microbial communities by improved fluorescence in situ hybridization techniques. Nat Rev Microbiol 6:339–348. doi:10.1038/nrmicro1888
Amann RI, Binder BJ, Olson RJ, Chisholm SW, Devereux R, Stahl DA (1990a) Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl Environ Microbiol 56:1919–1925
Amann RI, Krumholz L, Stahl DA (1990b) Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J Bacteriol 172:762–770
Apprill A, Marlow HQ, Martindale MQ, Rappé MS (2009) The onset of microbial associations in the coral Pocillopora meandrina. ISME J 3:685–699. doi:10.1038/ismej.2009.3
Ballesteros E (2006) Mediterranean coralligenous assemblages: a synthesis of present knowledge. Oceanogr Mar Biol 44:123–195
Bayer FM (1981) Key to the genera of Octocorallia exclusive of Pennatulacea (Coelenterata: Anthozoa), with diagnoses of new taxa. Proc Biol Soc Wash 94:902–947
Bayer T, Arif C, Ferrier-Pagès C, Zoccola D, Aranda M, Voolstra CR (2013a) Bacteria of the genus Endozoicomonas dominate the microbiome of the Mediterranean gorgonian coral Eunicella cavolini. Mar Ecol Prog Ser 479:75–84. doi:10.3354/meps10197
Bayer T, Neave MJ, Alsheikh-Hussain A, Aranda M, Yum LK, Mincer T, Hughen K, Apprill A, Voolstra CR (2013b) The microbiome of the Red Sea coral Stylophora pistillata is dominated by tissue-associated Endozoicomonas bacteria. Appl Environ Microbiol 79:4759–4762. doi:10.1128/AEM.00695-13
Bobrow MN, Harris TD, Shaughnessy KJ, Litt GJ (1989) Catalyzed reporter deposition, a novel method of signal amplification. Application to immunoassays. J Immunol Methods 125:279–285. doi:10.1016/0022-1759(89)90104-X
Bourne DG, Garren M, Work TM, Rosenberg E, Smith GW, Harvell CD (2009) Microbial disease and the coral holobiont. Trends Microbiol 17:554–562. doi:10.1016/j.tim.2009.09.004
Bright M, Bulgheresi S (2010) A complex journey: transmission of microbial symbionts. Nat Rev Microbiol 8:218–230. doi:10.1038/nrmicro2262
Burge CA, Mouchka ME, Harvell CD, Roberts S (2013) Immune response of the Caribbean sea fan, Gorgonia ventalina, exposed to an Aplanochytrium parasite as revealed by transcriptome sequencing. Front Physiol 4:180. doi:10.3389/fphys.2013.00180
Coma R, Gili JM, Zabala M, Riera T (1994) Feeding and prey capture cycles in the aposymbiotic gorgonian Paramuricea clavata. Mar Ecol Prog Ser 115:257–270. doi:10.3354/meps115257
Coma R, Ribes M, Zabala M, Giti JM (1995) Reproduction and cycle of gonadal development in the Mediterranean gorgonian Paramuricea clavata. Mar Ecol Prog Ser 117:173–183. doi:10.3354/meps117173
Coma R, Ribes M, Serrano E, Jiménez E, Salat J, Pascual J (2009) Global warming-enhanced stratification and mass mortality events in the Mediterranean. Proc Natl Acad Sci USA 106:6176–6181. doi:10.1073/pnas.0805801106
Daims H, Brühl A, Amann R, Schleifer KH, Wagner M (1999) The domain-specific probe EUB338 is insufficient for the detection of all bacteria: development and evaluation of a more comprehensive probe set. Syst Appl Microbiol 22:434–444. doi:10.1016/S0723-2020(99)80053-8
Ding JY, Shiu JH, Chen WM, Chiang YR, Tang SL (2016) Genomic insight into the host-endosymbiont relationship of Endozoicomonas montiporae Cl-33T with its coral host. Front Microbiol 7:251. doi:10.3389/fmicb.2016.00251
Dove SG, Hoegh-Guldberg O, Ranganathan S (2001) Major colour patterns of reef-building corals are due to a family of GFP-like proteins. Coral Reefs 19:197–204. doi:10.1007/PL00006956
Franzenburg S, Walter J, Künzel S, Wang J, Baines JF, Bosch TC, Fraune S (2013) Distinct antimicrobial peptide expression determines host species-specific bacterial associations. Proc Natl Acad Sci USA 110:E3730–E3738. doi:10.1073/pnas.1304960110
Galloway SB, Work TM, Bochsler VS, Harley RA, Kramarsky-Winter E, McLaughlin SM, Meteyer CU, Morado JF, Nicholson JH, Parnell PG, Peters EC, Reynolds TL, Rotstein DS, Sileo L, Woodley CM (2007) Coral disease and health workshop: coral histopathology II. NOAA technical memorandums nos NCCOS 56 and CRCP 4. National Oceanic and Atmospheric Administration, Silver Spring, MD
Garrity GM, Bell JA, Lilburn T (2005) Oceanospirillales ord. nov. In: Brenner DJ, Krieg NR, Staley JT, Garrity GM et al (eds) Bergey’s manual of systematic bacteriology, 2nd edn. Springer, New York, pp 270–323
Goffredi SK, Johnson SB, Vrijenhoek RC (2007) Genetic diversity and potential function of microbial symbionts associated with newly discovered species of Osedax polychaete worms. Appl Environ Microbiol 73:2314–2323. doi:10.1128/AEM.01986-06
Hansson L, Agis M, Maier C, Weinbauer MG (2009) Community composition of bacteria associated with cold-water coral Madrepora oculata: within and between colony variability. Mar Ecol Prog Ser 397:89–102. doi:10.3354/meps08429
Hughes TP, Graham NA, Jackson JB, Mumby PJ, Steneck RS (2010) Rising to the challenge of sustaining coral reef resilience. Trends Ecol Evol 25:633–642. doi:10.1016/j.tree.2010.07.011
Jensen S, Duperron S, Birkeland NK, Hovland M (2010) Intracellular Oceanospirillales bacteria inhabit gills of Acesta bivalves. FEMS Microbiol Ecol 74:523–533. doi:10.1111/j.1574-6941.2010.00981.x
Krediet CJ, Ritchie KB, Paul VJ, Teplitski M (2013) Coral-associated micro-organisms and their roles in promoting coral health and thwarting diseases. Proc Biol Sci 280:20122328. doi:10.1098/rspb.2012.2328
Kvennefors EC, Leggat W, Hoegh-Guldberg O, Degnan BM, Barnes AC (2008) An ancient and variable mannose-binding lectin from the coral Acropora millepora binds both pathogens and symbionts. Dev Comp Immunol 32:1582–1592. doi:10.1016/j.dci.2008.05.010
Kvennefors EC, Sampayo E, Ridgway T, Barnes AC, Hoegh-Guldberg O (2010) Bacterial communities of two ubiquitous Great Barrier Reef corals reveals both site- and species-specificity of common bacterial associates. PLoS ONE 5:e10401. doi:10.1371/journal.pone.0010401
La Rivière M, Roumagnac M, Garrabou J, Bally M (2013) Transient shifts in bacterial communities associated with the temperate gorgonian Paramuricea clavata in the Northwestern Mediterranean Sea. PLoS ONE 8:e57385. doi:10.1371/journal.pone.0057385
La Rivière M, Garrabou J, Bally M (2015) Evidence for host specificity among dominant bacterial symbionts in temperate gorgonian corals. Coral Reefs 34:1087–1098. doi:10.1007/s00338-015-1334-7
Lee ST, Davy SK, Tang SL, Fan TY, Kench PS (2015) Successive shifts in the microbial community of the surface mucus layer and tissues of the coral Acropora muricata under thermal stress. FEMS Microbiol Ecol. doi:10.1093/femsec/fiv142
Lewis JC, Barnowski TF, Telesnicki GJ (1992) Characteristics of carbonates of gorgonian axes (Coelenterata, Octocorallia). Biol Bull 183:278–296. doi:10.2307/1542215
Lonkar SS, Kedar GT, Kotangale JP, Kale MA (2011) Dyes and chemicals used in biomaterial study as stains for invertebrates. Int J Chem Res 2:22–25
McKinstry MJ, Chapman GB, Spoon DM, Peters EC (1989) The occurrence of bacterial colonies in the epidermis of the tentacles of the sea anemone Aiptasia pallida. Trans Am Microsc Soc 108:239–244
Meyer JL, Paul VJ, Teplitski M (2014) Community shifts in the surface microbiomes of the coral Porites astreoides with unusual lesions. PLoS ONE 9:e100316. doi:10.1371/journal.pone.0100316
Meziti A, Ramette A, Mente E, Kormas KA (2010) Temporal shifts of the Norway lobster (Nephrops norvegicus) gut bacterial communities. FEMS Microbiol Ecol 74:472–484. doi:10.1111/j.1574-6941.2010.00964.x
Morrow KM, Moss AG, Chadwick NE, Liles MR (2012) Bacterial associates of two Caribbean coral species reveal species-specific distribution and geographic variability. Appl Environ Microbiol 78:6438–6449. doi:10.1128/AEM.01162-12
Mouchka ME, Hewson I, Harvell CD (2010) Coral-associated bacterial assemblages: current knowledge and the potential for climate-driven impacts. Integr Comp Biol 50:662–674. doi:10.1093/icb/icq061
Moya A, Peretó J, Gil R, Latorre A (2008) Learning how to live together: genomic insights into prokaryote-animal symbioses. Nat Rev Genet 9:218–229. doi:10.1038/nrg2319
Neulinger SC, Gartner A, Jarnegren J, Ludvigsen M, Lochte K, Dullo W-C (2009) Tissue-associated “Candidatus Mycoplasma corallicola” and filamentous bacteria on the cold-water coral Lophelia pertusa (Scleractinia). Appl Environ Microbiol 75:1437–1444. doi:10.1128/AEM.01781-08
Nonaka M, Nakamura M, Tsukahara M, Reimer JD (2012) Histological examination of precious corals from the Ryukyu Archipelago. J Mar Biol 2012:1–14. doi:10.1155/2012/519091
Pernthaler A, Pernthaler J, Amann R (2002) Fluorescence in situ hybridization and catalyzed reporter deposition for the identification of marine bacteria. Appl Environ Microbiol 68:3094–3101. doi:10.1128/AEM.68.6.3094-3101.2002
Pett-Ridge J, Weber PK (2012) NanoSIP: NanoSIMS applications for microbial biology. Methods Mol Biol 881:375–408. doi:10.1007/978-1-61779-827-6_13
Raina JB, Tapiolas D, Willis BL, Bourne DG (2009) Coral-associated bacteria and their role in the biogeochemical cycling of sulfur. Appl Environ Microbiol 75:3492–3501. doi:10.1128/AEM.02567-08
Rosenberg E, Koren O, Reshef L, Efrony R, Zilber-Rosenberg I (2007) The role of microorganisms in coral health, disease and evolution. Nat Rev Microbiol 5:355–362. doi:10.1038/nrmicro1635
Rua CP, Trindade-Silva AE, Appolinario LR, Venas TM, Garcia GD, Carvalho LS, Lima A, Kruger R, Pereira RC, Berlinck RG, Valle RA, Thompson CC, Thompson F (2014) Diversity and antimicrobial potential of culturable heterotrophic bacteria associated with the endemic marine sponge Arenosclera brasiliensis. PeerJ 2:e419. doi:10.7717/peerj.419
Salih A, Larkum A, Cox G, Kühl M, Hoegh-Guldberg O (2000) Fluorescent pigments in corals are photoprotective. Nature 408:850–853. doi:10.1038/35048564
Schuett C, Doepke H, Grathoff A, Gedde M (2007) Bacterial aggregates in the tentacles of the sea anemone Metridium senile. Helgol Mar Res 61:211–216. doi:10.1007/s10152-007-0069-4
Sharp KH, Ritchie KB, Schupp PJ, Ritson-Williams R, Paul VJ (2010) Bacterial acquisition in juveniles of several broadcast spawning coral species. PLoS ONE 5:e10898. doi:10.1371/journal.pone.0010898
Speck MD, Donachie SP (2012) Widespread Oceanospirillaceae bacteria in Porites spp. J Mar Biol 2012:1–7. doi:10.1155/2012/746720
Sudek M, Work TM, Aeby GS, Davy SK (2012) Histological observations in the Hawaiian reef coral, Porites compressa, affected by Porites bleaching with tissue loss. J Invertebr Pathol 111:121–125. doi:10.1016/j.jip.2012.07.004
Tout J, Jeffries TC, Petrou K, Tyson GW, Webster NS, Garren M, Stocker R, Ralph PJ, Seymour JR (2015a) Chemotaxis by natural populations of coral reef bacteria. ISME J 9:1764–1777. doi:10.1038/ismej.2014.261
Tout J, Siboni N, Messer LF, Garren M, Stocker R, Webster NS, Ralph PJ, Seymour JR (2015b) Increased seawater temperature increases the abundance and alters the structure of natural vibrio populations associated with the coral Pocillopora damicornis. Front Microbiol 6:432. doi:10.3389/fmicb.2015.00432
Wallner G, Amann R, Beisker W (1993) Optimizing fluorescent in situ hybridization with rRNA-targeted oligonucleotide probes for flow cytometric identification of microorganisms. Cytometry 14:136–143. doi:10.1002/cyto.990140205
Webster NS, Wilson KJ, Blackall LL, Hill RT (2001) Phylogenetic diversity of bacteria associated with the marine sponge Rhopaloeides odorabile. Appl Environ Microbiol 67:434–444. doi:10.1128/AEM.67.1.434-444.2001
Wood-Charlson EM, Hollingsworth LL, Krupp DA, Weis VM (2006) Lectin/glycan interactions play a role in recognition in a coral/dinoflagellate symbiosis. Cell Microbiol 8:1985–1993. doi:10.1111/j.1462-5822.2006.00765.x
Work TM, Aeby GS (2014) Microbial aggregates within tissues infect a diversity of corals throughout the Indo-Pacific. Mar Ecol Prog Ser 500:1–9. doi:10.3354/meps10698
Acknowledgments
The authors would like to thank Mathieu Fallet, Fanny Baudimont and Lionel Chasson (Centre d’Immunologie de Marseille-Luminy) for their assistance with the preparation of the histological samples and confocal microscopy. We are grateful to Christian Tamburini (Mediterranean Institute of Oceanography) for providing access to the laboratory facilities used during this study. We also thank Esther Peters (George Mason University) for her valuable guidance throughout the analysis of histological sections and Joaquim Garrabou (Institut de Ciencies del Mar—CSIC) for his constant interest and encouragement, and Chantal Bézac, Jean-Baptiste Raina and Marc Tedetti for improving the FISH protocol. Finally, we thank the scientific divers of the OSU Pythéas Institute for their assistance with the sample collection.
Funding
This study was funded by the Total Foundation for Marine Biodiversity (Grant Number DN2024Y9TOTAL).
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La Rivière, M., Garel, M. & Bally, M. Localization of endobacteria in the gastrodermis of a Mediterranean gorgonian coral, Paramuricea clavata, using fluorescence in situ hybridization. Mar Biol 163, 206 (2016). https://doi.org/10.1007/s00227-016-2975-y
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DOI: https://doi.org/10.1007/s00227-016-2975-y