Phylogenetic survey of metabolically active microbial communities associated with the deep-sea coral Lophelia pertusa from the Apulian plateau, Central Mediterranean Sea

ARTICLE IN PRESS

Deep-Sea Research I 53 (2006) 62–75 www.elsevier.com/locate/dsr

Phylogenetic survey of metabolically active microbial communities associated with the deep-sea coral Lophelia pertusa from the Apulian plateau, Central Mediterranean Sea Michail M. Yakimova,, Simone Cappelloa, Ermanno Crisafia, Angelo Tursib, Alessandra Savinic, Cesare Corsellic, Simona Scarfid, Laura Giulianoa a

Istituto per l’Ambiente Marino Costiero (IAMC) sez. Messina, Istituto Sperimentale Talassografico (IST) Messina, Spianata S. Raineri, 86 I-98122 Messina, Italy b Department of Zoology, University of Bari, Italy c Department of Geological Sciences and Geotechnologies, Milano-Bicocca University, Milan, Italy d Department of Animal Biology and Marine Ecology, Messina University, Messina, Italy Received 31 January 2005; received in revised form 4 July 2005; accepted 31 August 2005 Available online 2 November 2005

Abstract Living deep-water coral assemblages were discovered recently inhabiting the Mediterranean Sea between the depths of 300 and 1000 m off the Cape of Santa Maria di Leuca (Apulian platform, Ionian Sea). This living assemblage was dominated by two colonial scleractinian corals, Lophelia pertusa and Madrepora oculata. Two other corals, Desmophyllum crystagalli and Stenocyathus vermiformis were also recovered from this site, but were much less common. The composition of the metabolically active fraction of the microbial community associated with living specimens of L. pertusa was determined. Dead corals, proximal sediments and overlying seawater were also sampled and analyzed. Complementary 16S ribosomal DNA (crDNA) was obtained from total RNA extracted from all samples that had been subjected to reverse transcription-PCR amplification. Domain-specific 16S PCR primers were used to construct four different 16S crDNA libraries containing 45 Archaea and 201 Bacteria clones. Using Archaea-specific primers, no amplification products were obtained from any coral samples (live and dead). Living specimens of L. pertusa seem to possess a specific microbial community different from that of dead coral and sediment samples. The majority of all coral-associated riboclones was related to the Holophaga-Acidobacterium and Nitrospira divisions (80%). Moreover, more than 12% of all coral-associated riboclones formed a separate deep-branching cluster within the a-Proteobacteria with no known close relatives. The metabolically active fraction of the bacterial community colonizing the dead corals was dominated by Proteobacteria related to the gamma and epsilon subdivisions (74% and 26% of all clones, respectively). Phylogenetic analysis of the Archaea clone library retrieved from proximal sediments indicated an exclusive dominance by the members of Crenarchaea Marine Group I (MGI), a lineage of unculturable microorganisms, widely distributed in marine habitats. In contrast, bacterial diversity was considerably higher in this sample than archaeal diversity, with four abundant eubacterial phylotypes: Proteobacteria, Verrucomicrobia, Actinobacteria and Planctomycetes (95% of all clones analyzed).

Corresponding author. Tel.: +39 090 66 9003; fax: +39 090 66 9007.

E-mail address: [email protected] (M.M. Yakimov). 0967-0637/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.dsr.2005.07.005

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

63

This is one of the first phylogenetic evaluation of the presumed metabolically active microbial community structure associated with the deep-sea scleratinian coral L. pertusa. r 2005 Elsevier Ltd. All rights reserved. Keywords: Mediterranean sea; Ionian sea; Apulian plateau; Deep-sea corals; Lophelia pertusa; Nitrospira; Holophaga; 16s crdna phylogenetic analysis; Microbial diversity; Coral-associated microbial community

1. Introduction Deep-sea coral reefs have been recorded from several areas of the Atlantic Ocean and to a lesser extent, the Mediterranean Sea (Le Danois, 1948; Pe´re`s and Picard, 1964; Zibrowius, 1980; Fossa˚ et al., 2002; Høvland et al., 2002). These communities are often dominated by the colonial scleratinian corals Lophelia pertusa and Madrepora oculata with which the solitary Desmophyllum cristagalli is generally associated (Le Danois, 1948; Pe´re`s and Picard, 1964). Most deep-water scleractinians collected from the Mediterranean are dead and covered by Fe–Mn oxides. Living corals have only seldomly been recovered. Delibrias and Taviani (1985) have suggested that the rarity of these coral banks in the Mediterranean was due to the homeothermic conditions in deep waters (12.7–14.5 1C, salinity 38.4–39.0%). Coral reefs represent some of the most biodiverse marine habitats providing ecological niches and substrates for a multitude of other eukaryotic species (Rogers, 1999). From recent studies of shallow water coral microbial ecology, it is known that coral-associated microbes are distinct from those in the water column, and that there appear to be coral species-specific microbial communities (Santavy et al., 1995; Kuhl et al., 1995; Rohwer et al. 2001, 2002; Knowlton and Rohwer, 2003). Moreover, coral-associated microflora obviously represent one of the most complex and important components of the biodiversity of coral communities (Rohwer et al., 2001; Frias-Lopez et al. 2002). Most of the associated Bacteria and Archaea have found to be represented by novel species, not present in the overlying seawater (Kellogg, 2004; Wegley et al., 2004). Several publications have attributed the coral diseases to the bacterial infections and reported a difference in the bacterial community composition between bleached and normal coral (Bythell et al., 2002; Cooney et al., 2002; Rosenberg and Ben-Haim, 2002; Frias-Lopez et al., 2004b). Differences in the bacterial community are apparent between healthy and diseased

corals, and even more telling, between diseased and apparently healthy tissues distant from disease lesions (Pantos et al. 2003; Friaz-Lopez et al., 2004a). However, recent studies of Cansas et al. (2004) raised doubts about this observation as no dramatic changes in the composition of the microbial community associated with white band disease type I of Acroporid coral populations were found. Moreover, the authors proposed that still unknown, non-bacterial pathogen might be the cause of this disease. The diversity of microbial associates likely has important evolutionary and ecological implications. However, very little is known about the type of interaction between the corals and microbes: is it merely a passive opportunistic association with Bacteria from the overlying seawater column, or are there specific commensal associations? These microbes may be involved in at least four different processes: (i) nutrient acquisition, i.e., cycling carbon, chelating iron, transforming molecular nitrogen, nitrate or polyphosphate into bioavailable form for corals (Vacelet et al., 1995; Wilkinson et al., 1992; Williams et al., 1987); (ii) processing of metabolic waste by eliminating it via the biotransformation of several toxic metabolites (Ritchie and Smith, 1995); (iii) stabilization and development of the skeleton by induction of calcareous settlement and triggering the metamorphosis of scleratinian corals (Webster et al., 2004); (iv) production of special secondary metabolites, such as protective antibiotics and other biologically active compounds (Bewley et al., 1996; Schmidt et al., 2000; Unson et al., 1994). In the case of deep-sea corals, the ecology is fundamentally different due to the lack of light and algal symbionts, cold temperature and elevated pressure. Consequentially, the coral-associated microbiology is likely to be unusual, reflecting the specific adaptation to thrive in such environment. The detailed investigation of commensal microflora is necessary for better understanding the overall biology of the organisms and the ecology of deep reefs, but up to now, this area is practically

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

64

unexplored and very few reports are available (Kellogg and Stone, 2004). The purpose of the present study was to perform the culture-independent 16S rRNA phylogenetic survey of the metabolically active microbial communities inhabiting living and dead L. pertusa corals and the surrounding environment. The coral reef chosen for analysis was on the deep-sea reef tract off the Cape of Santa Maria di Leuca, Apulian platform, Ionian Sea at a depth between 800 and 1000 m (Fig. 1). The following two fundamental questions have been addressed: (i) is there a coralspecific microbial population that is different from that of the prokaryotic community of the surrounding environments; and (ii) is there a distinction between microbial communities of the living and dead corals? Ribosomal RNA (rRNA) was used for our analysis as an indicator of metabolically active microbial populations, as rRNA is a highly labile molecule with a much shorter half-life than DNA and therefore should provide more accurate indicators of physiological status than DNA-based methods (Keer and Birch, 2003). Our results indicate that living coral tissue of L. pertusa is colonized by a specific microbial community that is distinct from that found in the surrounding environment and dead coral samples. This finding could support the hypothesis that deepsea scleratinian corals harbor a highly specialized microflora with which they interact symbiotically.

25m 50m 100m

41°N

250m 500m 750m 40°N 1000m +

1500m 2000m

39°N

2500m

2. Location, sampling and experimental 2.1. Study area and sample collections Living and dead L. pertusa specimens were harvested during the CORAL 2 cruise on board the R/V ‘‘Urania’’ at station 24 (39128.790 N, 18122.660 E) in August of 2002 (Tursi et al. 2004). The sampling area was about 20–25 miles off the Cape of Santa Maria di Leuca (LE), southern Italy (Fig. 1). All coral specimens (three living and four dead) used throughout this study were collected at a depth of 784 m using a sampling device consisting of an iron bar (1 m long, 60 cm diameter) with pieces of old fishing net attached (Tursi et al. 2004). The superficial sediments were sampled from the same trap. Coral samples were immediately placed in a sterile 50 ml Falcon tube, which was then tightly closed and immediately transported to the onboard laboratory. Each coral sample was further washed four times with 0.2 mm-filtered autoclaved seawater to remove any loosely associated microbes. To preserve RNA from degradation, coral samples were collected by removing a 2 cm by 1 cm portion and immersing it in 5 ml of RNAlaters solution (Ambion). Three superficial sediment subsamples (each 2 g) were directly treated with the same volume of RNAlaters solution. In order to compare the obtained diversity with that of the surrounding areas not containing deep-sea coral reefs, the sediment samples from reference sea bed site located 300 km to the south (36130.000 N, 15150.000 E) were sampled and analyzed in the same way. Immediately upon the return to shore, the RNAlaters solution within each tube was decanted and coral samples were crushed and homogenized in the tube, creating a slurry of coral tissue, mucus, microorganisms, and skeletal material. To determine if Bacteria in the water column were contaminating the coral samples, a 5 l water sample was taken from the water column immediately above the same colony. The water sample was recovered using a sampling rosette equipped with Niskin bottles (General Oceanics, Miami, Fl).

3000m 4000m 38°N 15°E

16°E

17°E

18°E

19°E

20°E

Fig. 1. Location of the deep-sea L. pertusa coral reefs. The map was constructed using Ocean Data View software (Schlitzer R., http://www.awi-bremenhaven.de/GEO/ODV, 2003). The sampling site is shown by a cross.

2.2. RNA extraction from coral and sediment samples and synthesis of 16S crDNA by reverse transcription reaction Extraction of RNA from corals and superficial sediment samples was performed using a DNA/ RNA extraction kit (QIAGEN, Valencia, CA)

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

65

according to the manufacturer’s protocol. Total RNA was precipitated with isopropanol, washed with 70% ethanol and after air-drying resuspended in 50 ml of diethylpyrocarbonate (DEPC)-treated sterile water. The quality of RNA samples was examined by agarose electrophoresis and concentrations determined using spectrophotometry (Biophotometer, Eppendorf). To increase the possibility of getting a positive amplification, samples from each type of material (living corals, dead corals, sediments), were pooled and subsequently purified into RNA-only aliquots by the DNase I treatment described elsewhere (Mills et al., 2004). The rRNAcrDNA heteroduplex was synthesized by reverse transcription using the 16S universal reverse primer Uni_1492R (50 -TACGYTACCTTGTTACGACTT30 ) (Lane, 1991) and SuperScript II RNase H-free reverse transcriptase (Life Technologies) according to the manufacturer’s protocol. Total RNA was initially denatured by heating at 70 1C for 10 min. The reverse transcription reaction mix consisted of 5 mM of a 16S rRNA reverse primer 1492R amplifying the vast majority of prokaryotic organisms, including both Bacteria and Archaea domains (Yakimov et al., 2001), 50–100 ng of denatured RNA, and 200 mM of deoxynucleoside triphosphate mix. The mixture was incubated for 5 min at 65 1C and 2 min at 4 1C, followed by the addition of 1
first-strand buffer (50 mM Tris–HCl [pH 8.3], 75 mM KCl, 3 mM MgCl2) and 75-U of RNase inhibitor and heating at 37 1C for 2 min. A 200U aliquot of SuperScript II RNaseH-free Reverse Transcriptase (Life Technologies) was added prior to a 50 min incubation at 42 1C that resulted in the transcription of RNA into complementary 16S ribosomal DNA (crDNA). The RT reaction was then stopped by heating the solution at 80 1C for 5 min, and the crDNA end product was used as the template for a standard PCR. Possible DNA contamination of RNA templates was routinely monitored by PCR amplification of aliquots of RNA that were not reverse transcribed. No contaminating DNA was detected in any of these reactions.

(Lane, 1991), and Archaea, Arc20_F (50 TTCCGGTTGATCCYGCCRG-30 ) (Hallam et al., 2003), primers. The PCR mix contained 10–50 ng of crDNA, 1
Qiagen reaction buffer, 1
solution Q (Qiagen), 1 pM of each forward and reverse primer, 200 mM dNTPs (Gibco), and 2.5 U of Qiagen Taq Polymerase. The PCR amplification involved a 3 min activation of the polymerase at 95 1C prior to 30 cycles each consisting of 1 min at 94 1C, 1 min at 50 1C and 2 min at 72 1C after which a 10 min extension at 72 1C was performed. PCR products were purified with QIAQuick PCR purification columns (Qiagen) and the amplicons were analyzed on 1.0% agarose gels run in Tris-borate-EDTA buffer stained with ethidium bromide and UV illuminated. Purified amplicons representing 16S crDNA sequences were cloned into the pGEM T-easy Vector II according to the manufacturer’s instructions (Promega). Inserts were subsequently PCR amplified from lysed colonies with primers specific for the vector, M13F (50 -GTAAAAC GACGGCCAG-30 ) and M13R (50 -CAGGAAA CAGCTATGAC-30 ). PCR products were digested (2 h, 37 1C) with MspI and HhaI for bacterial clones and with HhaI and RsaI for archaeal clones. Clones were grouped according to restriction fragment length polymorphism (RFLP) banding patterns, and representative clones were sequenced as previously described (Yakimov et al., 2002). RFLP groups containing two or more members had representative clones sequenced. Multiple representative clones were sequenced from RFLP groups containing five or more members to verify group integrity. All singletons (clones from those RFLP groups containing a single member) were sequenced. All calculations were based upon the number of clones incorporated in RFLP groups that had representative clones sequenced. Sequencing was performed with a BigDye Terminator v3.1 cycle sequencing kit on an automated capillary sequencer (model 3100 Avant Genetic Analyzer; Applied Biosystems).

2.3. PCR amplification, cloning and sequencing of PCR products

Four taxon-specific 16S rDNA forward primers (Table 1) were designed using the results of sequencing the living L. pertusa prokaryotic clone library. Only variable regions specific for the analyzed clones (CSCor-02Eub, CSCor-04Eub, CSCor-21Eub and CSCor-31Eub) were taken into considerations. The primers were checked for

Primers used for standard PCR amplification included the above reverse primer (Uni_1492R) and 16S rDNA forward domain-specific Bacteria, Bac27_F (50 -AGAGTTTGATCCTGGCTCAG-30 )

2.4. PCR amplification with taxon-specific primers

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

66

Table 1 Taxon-specific 16S rDNA forward primers used for RT–PCR analysis with total RNA isolated from living L. pertusa and seawater overlying the coral colony Primer

Annealing sitea

Sequence (50 -30 )

No. of RDP matchesb

Taxonomical affiliation

Size, (bp)c

CSCor-02 CSCor-04 CSCor-21 CSCor-31

239-258 541-560 432-451 739-758

CGGCCTACCAAGGCTACGAC GGGGGGGCAAGCGTTGTTCG GTAGCCCCGGCTAACTACGT GGCTTCCTGGACTAATACTG

9 44 No match 2

Nitrospira group Holophaga/Acidobacteria Phylum a-Proteobacteria g-Proteobacteria Achromatium assemblage

1253 951 1060 756

a

E.coli numbering. Only absolute matches to the RDP entries (no mismatches, insertions and deletions) obtained by Probe Match program (version 2.1r3) available from RDP web site (http://35.8.164.52/cgis/probe_match.cgi?su=SSU) were considered. c Length of fragments expected was calculated using sequences of corresponding riboclones. b

specificity and only absolute matches to the RDP entries (no mismatches, insertions and deletions) obtained by the Probe Match program (version 2.1r3) available from the RDP web site (http:// 35.8.164.52/cgis/probe_match.cgi?su=SSU) were considered. The chosen primers were applied further in the RT–PCR reaction with original RNA aliquotes as described above and the amplicons were analyzed on 1.0% agarose gels run in Trisborate-EDTA buffer stained with ethidium bromide and UV illuminated. 2.5. Phylogenetic and rarefaction analysis Sequence analysis was preformed as previously described (Yakimov et al., 2004). The sequences of individual inserts were initially aligned with the program BLAST 2 Sequences (Altschul et al., 1997; Tatusova and Madden, 1999) available through the National Center for Biotechnology Information. Sequences from this study and reference matching sequences, as determined by BLAST analysis, were subsequently aligned with the RDP Phylip Interface package (Maidak et al., 1997). Sequence data were checked using the CHECK_CHIMERA programme (Maidak et al., 1997) to detemine the presence of any hybrid sequences. Nucleotide sequences were manually aligned to 16S rRNA sequence data from the RDP database considering their secondary structure using the Se-Al sequence alignment editor version 1.0 alpha 1 (Rambaut, 1996). Further phylogenetic analyses were restricted to nucleotide positions that were unambiguously alignable in all sequences (600 nucleotides in average). Multiple bootstrapped data sets (1000 samplings) of the alignment data were exported as a

PHYLIP 3.5 interleaved file type to run the SEQBOOT program. Least-squares distance matrix analyses, based on evolutionary distances, were estimated from similarity values using the maximum likelihood analysis with multiple data sets option. Phylogenetic analyses using neighbor joining NEIGHBOR and parsimony DNAPARS methods were performed. Random order input of sequences, single jumbling and the global rearrangement option were used to avoid potential bias introduced by the order of sequence addition. The resulting tree files were analyzed using the CONSENSE program to provide confidence estimates for phylogenetic tree topologies and to make a majority rule consensus tree. All mentioned phylogenetic programs used were from the Phylogenetic Inference Package (PHYLIP) version 3.6 (Felsenstein, 2001). Rarefaction analysis was performed with the equations as described in Heck et al. (1975). Standard calculations were used to produce the curve with the total number of clones obtained compared to the number of clones representing each unique RFLP pattern. Regressions were performed using the algorithm for an exponential rise to a maximum: y ¼ a
ð1  ebx Þ. The asymptote of the curve (a) describes the estimated total diversity of the clone library. These values were used to estimate the percentage of total diversity sampled for each clone library. 2.6. Nucleotide sequence accession numbers The 37 bacterial and 41 archaeal 16S crDNA gene sequences have been deposited in the EMBL/ GenBank/DDBJ database under accession numbers AJ876924-AJ877001. A partial sequence of the 18S

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

rRNA gene of L. pertusa has also been deposited in the EMBL/GenBank/DDBJ under accession number AJ877002.

samples exhibited the lowest diversity as revealed by RFLP analysis. Forty-three clones were analyzed and five major OTUs were observed by RFLP profiling. Thirty-one (18 and 13) and 11 (8 and 3) clones were clustering in two large subgroups with matching RFLP patterns. Five representative clones were sequenced from each of these RFLP groups to verify group integrity. One clone occurred only once in the dead coral clone library. These patterns were unique and were not observed either in living L. pertusa or in sediment samples. Archaeal RT–PCR product was not detected from either coral sample. Of the 45 archaeal clones retrieved from sediment samples and screened by RFLP analysis, only two unique banding patterns were observed. Rarefaction analysis was performed for all libraries to determine the number of unique clones as a proportion of the estimated total diversity. In only two instances, namely archaeal sediment and dead coral clone library, did the rarefaction curves reach a clear saturation, indicating that further sampling of these clone libraries would not have revealed additional diversity (Fig. 2a, b). For the living L. pertusa library (Fig. 2c), the asymptote (a) ¼ 14.5 indicated that 83% of the estimated

3. Results 3.1. RFLP and rarefaction analyses A total of 201 bacterial and 45 archaeal clones containing 16S crDNA inserts, derived from both living and dead L. pertusa specimens and sediment samples have been de-replicated using restriction fragment length polymorphism (RFLP) analysis. On the basis of different bands profiling, 12 operative taxonomic units (OTUs) were observed for 86 (91.5%) sediment bacterial clones. RFLP patterns of remaining eight clones occurred only once in the sediment clone library. Sixty-four bacterial clones derived from living specimens of L. pertusa (CSCor clone library) had 12 separate banding patterns (OTUs). Almost 72% of the clones belonged to three major OTUs, while RFLP patterns of five clones were unique (each occurring only once in CSCor library). None of them was observed in the sediment samples. Dead coral

No. OTUs

5

10

(A)

4

8

3

6

2

4

1

2

0 0 20

10

20

30

40

50

0

16

20

12

15

8

10

4

5

0

(B)

0

25

(C)

67

10

20

30

40

(D)

0 0

10

20

30

40

50

60

70

0

10

20

30

40

50

60

70

80

90

No. 16S crDNA clones

Fig. 2. Rarefaction curves for coral L. pertusa and sediment-derived clone libraries. The expected number of RFLP patterns (OTUs), as determined by the analytical algorithm described in the experimental section, is plotted versus the number of clones sampled for dead and living L. pertusa specimens (B, C) and archaeal (A) and bacterial (D) clone libraries from sediments. The dotted lines represent 95% confidence intervals.

ARTICLE IN PRESS 68

M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

diversity was sampled. For the bacterial sediment clone library (Fig. 2d) the asymptote (a) ¼ 25.5 indicated that 82% of the estimated diversity was sampled. Although additional sampling of clones would be necessary to reveal the full extent of diversity in two last libraries, numerically dominant RFLP groups were obtained. Specifically, three dominant bacterial phylotypes represented by clones CSCor-02Eub, CSCor-04Eub and CSCor-30Eub from the living L. pertusa library comprised 25, 24 and 23% of all clones, respectively (Fig. 3). 3.2. Phylogenetic diversity of metabolically active Bacteria Fig. 3 provides a phylogenetic overview of eubacterial clones retrieved from corals and proximal sediments as determined by Neighbor-joining analysis of maximum likelihood values. Highdiversity assemblages of bacterial sequences were detected in sediments and living L. pertusa samples, but not in dead corals. A total of 37 partial sequences were identified: 20 bacterial sequences from sediments directly underlying the coral colonies and a total of 17 sequences from living (12 sequences) and dead (5 sequences) coral samples. All clones fell within 12 different bacterial divisions. Division-level microbial diversity in each sample was estimated by dividing the number of clones representing each division by the total number of clones in the libraries. The sediment sample clone library consistently exhibited high division level diversity (Figs. 3 and 4), with the most abundant sequences representing different subdivisions of Proteobacteria (53% of all clones). The next most abundant groups of clones represented the Verrucomicrobia (18%), Planctomycetes (16%) and Actinobacteria (11%) divisions. Single clone CSSed17Eub was affiliated with the unidentified bacterium NKB19, found in deep-sea sediments. The living L. pertusa tissue clone library contained a significantly different bacterial sequence assemblage. Only 19% of the sequences were represented by Proteobacteria in stark contrast to the dominance of this division in the underlying sediments (Figs. 3 and 4). 80% of all clones represented only two divisions, Holophaga-Acidobacteria and Nitrospira (55% and 25%, respectively). More than 12% of all coral-associated clones formed a separate deep-branching cluster within the a-Proteobacteria with no known close

relatives. Remarkably, none of these bacterial sequences were detected in any other libraries. The dead L. pertusa sample clone library contained the lowest sequence diversity, representing two phylogenetic lineages of Eubacteria. The clone library was dominated by g-Proteobacteria (75% of all clones), while the remaining clones fell into deeply branched group of unclassified Eubacteria, absent in the underlying sediments and live coral tissues. 3.3. Phylogenetic diversity of metabolically active Archaea No archaeal sequences were detected in either coral clone libraries (living and dead). A total of 45 rRNA-derived Archaea clones obtained from the underlying sediments and their phylogenetic affiliation were estimated (Fig. 5). Interestingly, all sequences fell into one phylogenetic lineage, Marine Group I (MGI) of Crenarchaeota. Although the riboclones belonging to MGI group from the coral sample were also found from the reference site not containing deep-sea coral reefs, this group was not dominant there and found alongside archaeal representatives from nine other divisions belonging to Crenarchaeota and Euryarchaeota classes (data not shown). 3.4. Specificity of microbial community associated with L. pertusa We developed a molecular tool for the specific detection of four major Eubacteria phylotypes observed in living L. pertusa tissues. Based on the 16S crDNA sequencing results of CSCor-21Eub (deep branching a-Proteobacteria), CSCor31-Eub (g-Proteobacterium, distantly related to L. floridana symbiont), CSCor-04Eub (Holophaga) and CSCor02Eub (Nitrospira) riboclones, four forward primers, specifically targeting the hypervariable regions of corresponding 16S crDNA, were designed in order to detect their presence in the surrounding environment. Only absolute matches to the RDP entries (no mismatches, insertions or deletions) obtained by the Probe Match program (version 2.1r3) available from RDP web site (http:// 35.8.164.52/cgis/probe_match.cgi?su=SSU) were considered. These primers were tested with the total RNA isolated from living L. pertusa tissues for amplification of taxon-specific sequence fragments. Fig. 6 shows that amplification products could be

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

69

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

70

obtained in three cases. We failed to get an amplification product with the CSCor31 primer, possibly due to either primer–primer interactions or annealing difficulties. The remaining designed primers were shown to be specific to their group, as only one amplification product of expected size was obtained in each case (Fig. 6). In addition, further sequencing of obtained products confirmed that these fragments corresponded to the riboclone sequences upon which the primers were based. To determine if the source of coral-associated Bacteria was the surrounding water, a 5-l seawater sample was collected from the water column immediately above the L. pertusa colony. Total RNA was isolated, reverse transcripted and amplified using taxon-specific primers designed for this study (Table 1). No PCR products were observed by

100 LC PS DC

90

20

Planctomycetes

NKB19 division

30

Nitrospirae

40

Verrucomicrobia

50

Unclassified Eubacteria

60

Actinobacteria

Relative abundance, %

70

Candidate division OP3

Holophaga .Acidbacteria

80

10 0

γ α δ Proteobacteria

Fig. 4. Distribution of major eubacterial phylogenetic groups recovered from 16S crDNA clone libraries obtained from living coral (LC), dead coral (DC) and proximal sediments (PS). The relative abundance was used to standardize the difference for the number of clones observed in different clone libraries.

applying all four primers (Fig. 6). Therefore the surrounding water, as in the case of the sediment sample, did not appear to be the source of coralassociated Bacteria belonged to Holophaga, Nitrospira and a-Proteobacteria divisions. 4. Discussion Deep-sea corals are slow-growing stony colonies that have adapted for life in cold, sub-photic zone waters. These corals lack the photosynthetic symbiotic algae found in the many colorful shallow water filter-feeding corals. Healthy and well developed Lophelia–Madrepora–Desmophillum banks have been recently identified in the Ionian sea off the Apulian platform between the depth of 300 to 1000 m. This area was previously characterized as exhibiting unique oceanographic conditions likely enhancing nutrient availability for deep-sea corals. Hydrography and current dynamics appear to play an important role in the formation of deep-sea coral banks in regions of strong currents and/or the development of internal waves and where sediment accumulation is low. Moreover, because some L. pertusa colonies occur in areas where hydrocarbons seep into the water column from the seabed, it has been proposed that chemotrophic Bacteria inhabit these sites providing the corals and other suspension feeders with substantial and reliable food sources as well as skeletal materials (Høvland and Thomsen, 1997; Høvland et al., 1998; Høvland and Risk, 2003). Numerous studies have examined interactions between shallow water corals and microbes (see Rohwer and Kelley, 2004; Ritchie and Smith, 2004 for further references). These studies have shown that there is a dynamic microbiota living on the mucopolysaccharide surface layers, and possibly within the tissue of corals and in the surrounding reef waters. However, it is still not known whether microbes play specific symbiotic roles in coral biology or if the observed associations are merely opportunistic interactions of the coral animal

Fig. 3. Rooted phylogenetic tree of eubacterial clones clustered by Neighbor-joining of maximum likelihood values showing affiliation of partial bacterial 16S rRNA gene sequences to closest related sequences from either cultivated or cloned members of different microbial clusters. Clones obtained in this work are indicated in bold-type. Clones retrieved from living and dead corals are highlighted in black and grey, respectively. The number of similar clones with 98% identity cutoff is shown in brackets beside each representative clone. Percentages of 1000 bootstrap resampling that supported the branching orders in each analysis are shown above or near the relevant nodes (only values X50% are shown). The tree was rooted with 16S rRNA gene sequences of Methanococcus jannaschii (M59126). The scale bar indicates 5% estimated sequence divergence.

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

71

Fig. 5. Rooted phylogenetic tree of archeal clones retrieved from coral-underlying sediments. The tree was constructed as described in Fig. 3 and rooted and outgrouped with 16S rRNA gene sequences of Psychroserpens burtonensis (U62913) and Methanococcus jannaschii (M59126), respectively.

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

72

A.

B.

C.

D.

A.

B.

C.

D. 1000 500

Fig. 6. PCR amplification using sequence-specific primers targeting hypervariable regions of 16S crDNA of: a-Proteobacterium riboclone CSCor-21EUB (A); Holophaga-Acidobacteriumrelated riboclone CSCor-04Eub (B); Nitrospira-related riboclone CSCor-02Eub (C); g-proteobacterium riboclone CSCor31-Eub (D). Results were obtained with total RNA isolated from both alive coral L.pertusa and ovelying seawater samples. The obtained RT–PCR products were displayed left and right from the molecular weight ladder, respectively.

colony with the water-column or sediment Bacteria (Ritchie and Smith, 2004). A few studies have suggested that corals may associate with specific microbes belonging to Eubacteria and Archaea kingdoms (Kellogg, 2004; Wegley et al., 2004). It has been shown that some harbor nitrogen-fixing Bacteria, suggesting that they may obtain fixed nitrogen from associated microbes that are fed in anaerobic environments within the colony (Sorokin, 1973; Shashar et al., 1994; Kuhl et al., 1995; Shashar et al., 1997). Corals may encourage the growth of microbes by secreting fixed carbon in the form of mucus and then feed upon them (Ritchie and Smith, 1995). Finally, specialized microbiota may be important for protecting the coral animal from pathogens by occupying entry niches and/or through the production of secondary metabolites (i.e. antibiotics) as demonstrated for some sponges (Schmidt et al., 2000; Unson et al., 1994). In order to study coral–microbial interactions, it is necessary to determine which microbes are actually living on or within corals. With the advent of culture-independent molecular techniques it is now possible to identify whole microbial diversity without the initial bias of culturing. Moreover, the RNA-based techniques should provide a more accurate survey of metabolically active microbial populations thriving in the studied environment (Keer and Birch, 2003). To our knowledge, the present study is one of the first attempts to characterize the microbial population associated with deep-sea corals (Kellogg and Stone, 2004). Comparing the microbial communities inhabiting the underlying sediments and the surrounding seawater with those directly retrieved

from the L. pertusa coral bodies, we found that several coral-associated riboclones were phylogenetically affiliated with bacterial divisions not present in the surrounding environments. This finding strongly suggests the existence of symbiotic relationships between deep-sea L. pertusa corals and the associated microbial population. The majority of all coral-derived sequences are related to the Holophaga-Acidobacterium division, suggesting a close relationship with the recently described sponge symbionts (Hentschel et al., 2002). The second major group of Bacteria found only in living corals represent the Nitrospira division, of which the members identified from our study are known nitrifiers, oxidizing nitrite to molecular nitrogen. This action could be very helpful in allowing the coral to eliminate toxic nitrite from the colony. Conspicuously, more than 12% of all coral-associated clones formed a separate deeply branching cluster within the a-Proteobacteria division with no known close relatives. The finding that L. pertusa may be involved in a specific association with aProteobacteria is not surprising as many Bacteria of this division are found as abundant, and potentially important components of the coral holobiont (Casas et al., 2004; Breitbart et al., 2005). Currently, we do not know the exact basis of this particular coral–microbe interaction, but the maintenance of this specific microbial population, distinct from that observed in surrounding environment, may be important for the thriving of L. pertusa in the deep sea. No archaeal sequences were obtained from the coral samples, while substantial numbers were recovered from the underlying sediments. This observation is in contrast with recent finding of coral-associated Archaea. Using culture-independent techniques, it was show that Archaea were abundant and widespread on corals. Sequence analyses of Archaea on three species of Caribbean corals revealed that coral-associated Archaea are novel, diverse, and include representatives from both the Crenarchaeota and Euryarchaeota (Kellogg, 2004; Wegley et al., 2004). The phylogenetic analysis of sediment archaeal populations revealed that all riboclones represented only the Crenarchaea MGI. The vast majority of organisms belonging to Crenarchaeota were isolated from marine environments, including areas where hydrocarbons seep into the water column from the sea-bed (Fuhrman and Davis, 1997). Since the majority of MGI organisms are uncultivable, at least for now, we

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

cannot speculate about their functional role in the deep-sea environment. But such a decrease of diversity, especially compared with that detected in ‘‘normal’’ deep-sea sediments where at least 15 different phylotypes were obtained, could indirectly indicate unique conditions in the studies area (LaMontagne et al., 2004). One of the possible explanations of such dominance of the MGI Archeae is that these organisms are somehow involved in the formation of chemolithotrophic communities driven by deep-sea cold hydrocarbon seepage. Based on our phylogenetic survey of metabolically active microbial communities associated with Lophelia pertusa found on Apulian Plateau, Central Mediterranean Sea, we can postulate that these deep-sea corals are harboring specific microbial populations. Due to several synergetic effects, these may allow the existence of these corals in such an extreme environment. Further investigations of coral–microbe associations are needed to understand the ecology of deep-sea corals. Acknowledgements We thank Carmen Fato for excellent technical assistance. We are indebted to the crew of R/V ‘‘Urania’’ and to the scientific team help with the collection and on-board treatment of the numerous samples. We would also like to thank Andrew Dalby for improving the manuscript. This work was funded by EC Project COMMODE EVK3-CT200200077 under the Framework V program.

References Altschul, S.F., Madden, T.L., Scha¨ffer, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSIBLAST: a new generation of protein database search programs. Nucleic Acids Research 25, 3389–3402. Bewley, C.A., Holland, N.D., Faulkner, D.J., 1996. Two classes of metabolites from Theonella swinhoei are localized in distinct populations of bacterial symbionts. Experientia 52, 716–722. Breitbart, M., Bhagooli, R., Griffin, S., Johnston, I., Rohwer, F., 2005. Microbial communities associated with skeletal tumors on Porites compressa. FEMS Microbiology Letters 243 (2), 431–436. Bythell, J.C., Barer, M.R., Cooney, R.P., Guest, J.R., O’Donnell, A.G., Pantos, O., Le Tissier, M.D., 2002. Histopathological methods for the investigation of microbial communities associated with disease lesions in reef corals. Letters to Applied Microbiology 34 (5), 359–364.

73

Casas, V., Kline, D.I., Wegley, L., Yu, Y., Breitbart, M., Rohwer, F., 2004. Widespread association of a Rickettsialeslike bacterium with reef-building corals. Environmental Microbiology 6 (11), 1137–1148. Cooney, R.P., Pantos, O., Le Tissier, M.D., Barer, M.R., O’Donnell, A.G., Bythell, J.C., 2002. Characterization of the bacterial consortium associated with black band disease in coral using molecular microbiological techniques. Environmental Microbiology 4 (7), 401–413. Delibrias, G., Taviani, M., 1985. Dating the death of Mediterranean deep-sea scleractinian corals. Marine Geology 62, 175–180. Felsenstein, J., 2001. PHYLIP phylogenetic inference package, version 3.6 ed. Department of Genetics, University of Washington. Seattle: WA, USA. Fossa˚, J.H., Mortensen, P.B., Furevik, D.M., 2002. The deepwater coral Lophelia pertusa in Norwegian waters: distribution and fishery impacts. Hydrobiologia 471, 1–12. Fuhrman, J.A., Davis, A.A., 1997. Widespread archeae and novel bacteria from the deep sea as shown by 16S rRNA gene sequences. Marine Ecology Progress Series 150, 275–285. Frias-Lopez, J., Zerkle, A.L., Bonheyo, G.T., Fouke, B.W., 2002. Partitioning of bacterial communities between seawater and healthy, black band diseased, and dead coral surfaces. Applied and Environmental Microbiology 68, 2214–2228. Frias-Lopez, J., Bonheyo, G.T., Fouke, B.W., 2004a. Identification of differential gene expression in bacteria associated with coral black band disease by using RNA-arbitrarily primed PCR. Applied and Environmental Microbiology 70 (6), 3687–3694. Frias-Lopez, J., Klaus, J.S., Bonheyo, G.T., Fouke, B.W., 2004b. Bacterial community associated with black band disease in corals. Applied and Environmental Microbiology 70 (10), 5955–5962. Hallam, S.J., Girguis, P.R., Preston, C.M., Richardson, P.M., DeLong, E.F., 2003. Identification of methyl coenzyme M reductase A (mcrA) genes associated with methane-oxidizing Archeae. Applied and Environmental Microbiology 69 (9), 5483–5491. Heck, K.L., Belle, G.V., Simberloff, D., 1975. Explicit calculation of the rarefaction diversity measurement and the determination of sufficient sample size. Ecology 56, 1459–1461. Hentschel, U., Hopke, J., Horn, M., Friedrich, A.B., Wagner, M., Hacker, J., 2002. Molecular evidence for a uniform microbial community in sponges from different oceans. Applied and Environmental Microbiology 68, 4431–4440. Høvland, M., Risk, M., 2003. Do Norvegian deep-water coral reefs rely on seeping fluids? Marine Geology 1–2, 83–96. Høvland, M., Thomsen, E., 1997. Cold-water corals—Are they hydrocarbon seep related? Marine Geology 137, 159–164. Høvland, M., Mortensen, P.B., Brattegard, T., Strass, P., Rokoengen, K., 1998. Ahermatypic coral banks off midNorway: evidence for a link with seepage of light hydrocarbons. Palaios 13, 189–200. Høvland, M., Vasshus, S., Indreeide, A., Austdal, L., Nilsen, Ø., 2002. Mapping and imaging deep-sea coral reefs off Norway, 1982–2000. Hydrobiologia 471, 13–17.

ARTICLE IN PRESS 74

M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75

Keer, J.T., Birch, L., 2003. Molecular methods for the assessment of bacterial viability. Journal of Microbiological Methods 53, 175–183. Kellogg, C.A., 2004. Tropical Archaea: diversity associated with the surface microlayer of corals. Marine Ecology Progress Series 273, 81–88. Kellogg, C.A., Stone, R.P., 2004. A pilot study of deep water coral microbial ecology. In: ASLO/TOS Ocean Research Conference, Honolulu, February, 15–20. Knowlton, N., Rohwer, F., 2003. Multispecies microbial mutualisms on coral reefs: the host as a habitat. American Naturalist 162 (4), 51–62. Kuhl, M., Cohen, Y., dalsgaard, T., Jorgenson, B.B., Revsbech, N.P., 1995. Microenvironment and photosynthesis of zzoxanthellae in scleratinian corals studied wth microsensors for O2, pH and light. Marine Ecology Progress Series 117, 159–172. LaMontagne, M.G., Leifer, I., Bergmann, S., Van De Werfhorst, L.C., Holden, P.A., 2004. Bacterial diversity in marine hydrocarbon seep sediments. Environmental Microbiology 6 (8), 799–808. Lane, D.J., 1991. 16S/23S rRNA Sequencing. In: Stackerbrandt, E., Goodfellow, M. (Eds.), Nucleic acid techniques in bacterial systematics. Wiley, New York, pp. 115–175. Le Danois, E., 1948. Les Profondeurs de la Mer. Payot, Paris, pp. 303. Maidak, B.L., Olsen, G.J., Larsen, N., Overbeek, R., McCaughey, M.J., Woese, C.R., 1997. The RDP (Ribosomal Database Project). Nucleic Acids Research 25, 109–111. Mills, H.J., Martinez, R.J., Story, S., Sobecky, P.A., 2004. Identification of members of the metabolically active microbial populations associated with Beggiatoa species mat communities from Gulf of Mexico cold-seep sediments. Applied and Environmental Microbiology 70 (9), 5447–5458. Pantos, O., Cooney, R.P., Le Tissier, M.D., Barer, M.R., O’Donnell, A.G., Bythell, J.C., 2003. The bacterial ecology of a plague-like disease affecting the Caribbean coral Montastrea annularis. Environmental Microbiology 5 (5), 370–382. Pe´re`s, J.M., Picard, J., 1964. Nouveau Manuel de Bionomie benthique de la mer Me0 diterrane0 e. Recueil des Travaux de la Station Marine d’Endoume, 31 (47), pp 137. Rambaut, A., 1996. Se-Al (Sequence Alignment Editor) version 1.0 alpha 1. WWW Page, http://evolve.zoo.ox.ac.uk/Se-Al/ Se-Al.html. Ritchie, K.B., Smith, G.W., 1995. Preferential carbon utilization by surface bacterial communities from water mass, normal, and white-band diseased Acropora cervicornis. Molecular Marine Biology and Biotechnology 4, 345–354. Ritchie, K.B., Smith, G.W., 2004. Microbial communities of coral surface mucopolysaccharide layers. In: Rosenberg, E., Loya, Y. (Eds.), Coral Health and Disease. Springer Verlag, Berlin/Heidelberg, pp. 259–263. Rohwer, F., Breitbart, M., Jara, J., Azam, F., Knowlton, N., 2001. Diversity of bacteria associated with the Caribbean coral Montastraea franksi. Coral Reefs 20, 85–95. Rohwer, F., Seguritan, V., Azam, F., Knowlton, N., 2002. Diversity and distribution of coral-associated bacteria. Marine Ecology Progress Series 243, 1–10. Rohwer, F., Kelley, S., 2004. Culture-independent analyses of coral-associated microbes. In: Rosenberg, E., Loya, Y. (Eds.),

Coral Health and Disease. Springer Verlag, Berlin/Heidelberg, pp. 265–278. Rogers, A.D., 1999. The biology of Lophelia pertusa (Linnaeus 1758) and other deep-water reef-forming corals and impacts from human activities. International Review Hydrobiology: A Journal Covering all Aspects of Limnology and Marine Biology 84 (4), 315–406. Rosenberg, E., Ben-Haim, Y., 2002. Microbial diseases of corals and global warming. Environmental Microbiology 4 (6), 318–326. Santavy, D.L., Peters, E.C., Kozlowsky, J., Wilkinson, S., 1995. Characterization of the bacterium suspected in the incidence of white band disease. In: 95th General Meeting of the American Society of Microbiology, Washington DC, May 21–25, 1995. ASM, Washington. Schmidt, E.W., Obraztova, A.Y., Davidson, S.K., Faulkner, D.J., Haygood, M.G., 2000. Identification of the antifungal peptide-containing symbiont of the marine sponge Theonella swinhoei as a novel Delta-Proteobacterium Candidatus Entotheonella palauensis. Marine Biology 236, 969–977. Shashar, N., Cohen, Y., Loya, Y., Sar, N., 1994. Nitrogen fixation (acetylene reduction) in stony corals: evidence for coral-bacteria interactions. Marine Ecology Progress Series 111, 259–264. Shashar, N., Banaszak, A.T., Lesser, M.P., Amrami, D., 1997. Coral endolithic algae: life in a protected environment. Pacific Sciences 51, 167–173. Sorokin, Y.I., 1973. Trophical role of bacteria in the ecosystem of the coral reef. Nature 242, 415–417. Tatusova, T.A., Madden, T.L., 1999. BLAST 2 SEQUENCES, a new tool for comparing protein and nucleotide sequences. FEMS Microbiology Letters 177, 187–188. Tursi, A., Mastrototaro, F., Matarrese, A., Maiorano, P., D’Onghia, G., 2004. Biodiversity of the white coral reefs in the Ionian Sea (Central Mediterranean). Chemistry and Ecology 20 (Suppl. 1), 107–116. Unson, M.D., Holland, D., Faulkner, D.J., 1994. A brominated secondary metabolite synthesized by the cyanobacterial symbiont of a marine sponge and accumulation of the crystalline metabolite in the sponge tissue. Marine Biology 119, 1–11. Vacelet, J., Boury-Esnault, N., Fiale-Medion, A., Fischer, C.R., 1995. A methanotrophic carnivourous sponge. Nature 377, 269. Webster, N.S., Smith, L.D., Heyward, A.J., Watts, J.E., Webb, R.I., Blackall, L.L., Negri, A.P., 2004. Metamorphosis of a scleratinian coral in response to microbial biofilms. Applied and Environmental Microbiology 70 (2), 1213–1221. Wegley, L., Yu, Y., Breitbart, M., Casas, V., Kline, D.I., Rohwer, F., 2004. Coral-associated Archaea. Marine Ecology Progress Series 273, 89–96. Wilkinson, C.R., 1992. Symbiotic interaction between marine sponges and algae. In: Reisser, W. (Ed.), Algae and symbioses. Biopress, Bristol, England, pp. 112–151. Williams, W.M., Viner, A.B., Broughton, W.J., 1987. Nitrogen fixation (acetylene reduction) associated with the living coral Acropora variabilis. Marine Biology 94, 531–535. Yakimov, M.M., Giuliano, L., Timmis, K.N., Golyshin, P.N., 2001. Upstream-independent ribosomal RNA amplification analysis (URA): a new approach to characterizing the

ARTICLE IN PRESS M.M. Yakimov et al. / Deep-Sea Research I 53 (2006) 62–75 diversity of natural microbial communities. Environmental Microbiology 3 (10), 662–666. Yakimov, M.M., Giuliano, L., Crisafi, E., Chernikova, T.N., Timmis, K.N., Golyshin, P.N., 2002. Microbial community of a saline mud volcano at San Biagio-Belpasso, Mt. Etna (Italy). Environmental Microbiology 4 (5), 249–256. Yakimov, M.M., Gentile, G., Bruni, V., Cappello, S., D’Auria, G., Golyshin, P.N., Giuliano, L., 2004. Crude oil-induced

75

structural shift of coastal bacterial communities of Rod Bay (Terra Nova Bay, Ross Sea, Antarctica) and characterization of cultured cold-adapted hydrocarbonoclastic bacteria. FEMS Microbiology Ecology 49 (3), 419–432. Zibrowius, H., 1980. Les Scle´ractiniares de la Me´diterranee´ et de l’Atlantique nord-oriental. Memoires de Institut Oceanographique, Monaco 11, pp.107.