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An Improved Detection and Quantification Method for the Coral Pathogen Vibrio coralliilyticus

  • Bryan Wilson,

    Affiliation Centre for Marine Microbiology and Genetics, Australian Institute of Marine Science (AIMS), Townsville, Queensland, Australia

  • Andrew Muirhead,

    Affiliation Centre for Marine Microbiology and Genetics, Australian Institute of Marine Science (AIMS), Townsville, Queensland, Australia

  • Monika Bazanella,

    Affiliations Centre for Marine Microbiology and Genetics, Australian Institute of Marine Science (AIMS), Townsville, Queensland, Australia, Fachhochschule Krems, Krems, Austria

  • Carla Huete-Stauffer,

    Affiliations Centre for Marine Microbiology and Genetics, Australian Institute of Marine Science (AIMS), Townsville, Queensland, Australia, Università degli Studi di Genova, Genoa, Italy

  • Luigi Vezzulli,

    Affiliation Università degli Studi di Genova, Genoa, Italy

  • David G. Bourne

    Affiliation Centre for Marine Microbiology and Genetics, Australian Institute of Marine Science (AIMS), Townsville, Queensland, Australia

An Improved Detection and Quantification Method for the Coral Pathogen Vibrio coralliilyticus

  • Bryan Wilson, 
  • Andrew Muirhead, 
  • Monika Bazanella, 
  • Carla Huete-Stauffer, 
  • Luigi Vezzulli, 
  • David G. Bourne


DNA- and RNA-based PCR and reverse-transcription real-time PCR assays were developed for diagnostic detection of the vcpA zinc-metalloprotease implicated in the virulence of the coral pathogen Vibrio coralliilyticus. Both PCR methods were highly specific for V. coralliilyticus and failed to amplify strains of closely-related Vibrio species. The assays correctly detected all globally occurring V. coralliilyticus isolates including a newly-described isolate [TAV24] infecting gorgonians in the Mediterranean Sea and highlighted those isolates that had been potentially misidentified, in particular V. tubiashii strains ATCC 19105 and RE22, historically described as important oyster pathogens. The real-time assay is sensitive, detecting 10 gene copies and the relationships between gene copy number and cycle threshold (CT) were highly linear (R2≥99.7). The real-time assay was also not affected by interference from non-target DNA. These assays are useful for rapid detection of V. coralliilyticus and monitoring of virulence levels in environmental samples, allowing for implementation of timely management steps to limit and possibly prevent losses due to V. coralliilyticus infection, as well as furthering investigations of factors affecting pathogenesis of this important marine pathogen.


Coral reefs represent one of the most biologically diverse ecosystems in the world, as well as playing a vital role in supporting local communities by way of coastal protection, food production and tourism [1][3]. In recent decades however, coral reef health has been increasingly compromised by natural and anthropogenic disturbances [2] and reports of disease outbreaks have escalated [4]. Bacterial pathogens have been identified as causative agents for a number of these diseases [3] and in particular, members of the Vibrionaceae family [5][7] implicated in outbreaks. Of these, V. coralliilyticus is amongst the best studied [5], [8][21] and investigations into strains isolated from white syndrome disease outbreaks in the Indo-Pacific [6] revealed the expression of a zinc-metalloprotease demonstrated to cause coral tissue damage [15].

Zinc-metalloproteases are common in prokaryotic organisms, having an essential role in maintenance of cellular homeostasis. In pathogenic organisms (particularly opportunists), the action of these proteases enhances vascular permeability, necrotic tissue damage and cytotoxicity, therefore facilitating bacterial invasion [22] and the infection process. Expression of the metalloproteases in Vibrio species is growth-phase dependent and positively regulated by various quorum-sensing systems [23][27]. The zinc-metalloprotease of V. coralliilyticus has been shown to cause coral tissue lesions [8], [15] concurrent with white syndrome aetiology and so detection of this important virulence factor could provide early indications of infection and assist agencies in developing strategies to effectively contain coral disease outbreaks. There is a pressing need for disease diagnostics in coral science [28], as the principal method for coral disease detection currently involves field-based observations of gross macroscopic symptoms which, whilst requiring anatomical knowledge for correct diagnosis [29], also detect only the latest stages of disease, by which time remedial management measures are ineffective. PCR-based diagnostic methods are rapid, specific and sensitive with the detection of Vibrio species by these means previously validated in other systems [17], [30][34]. A real-time assay to detect V. coralliilyticus previously published targeted the dnaJ gene using the TaqMan qPCR system for amplification from cells and genomic DNA [17]. In this study, we describe the development of qualititative PCR and quantitative reverse transcription real-time PCR (using SYBR Green I dye) assays with an improved capacity not only for strain classification but also for quantification of viable and active V. coralliilyticus populations.


Standard curves and detection range

Standard curves constructed from serial tenfold dilutions of known concentrations (100–107 copies) of the pCR2.1-TOPO plasmid carrying the vcpA gene were successfully generated and allowed quantitative detection of the vcpA gene using both developed primer sets (vcpAF-vcpAR and vcpARTF-vcpARTR). Standard curves for both real-time PCR and PCR showed high linear correlation, with linear regression coefficients (R2 value) equal to or greater than 0.997 and both primer sets were able to detect the vcpA gene down to levels of 10 gene copies.

Evaluation of primers

In silico comparisons of the primer pairs against sequences stored in the NCBI database using BLASTn [35] revealed that both VcpAF and VcpARTF forward primers matched the V. tubiashii zinc-metalloprotease vtpA (Accession Numbers EU675309 and FJ455119-FJ455121) though neither of the reverse primers (VcpAR and VcpARTR) matched any non-target species. Using a standard PCR approach with the primer set VcpAF and VcpAR, a single amplicon (1.8 kB) was observed when DNA from all globally-occurring V. coralliilyticus strains (Table 1) was added to single reactions, the exception being strain P3 (LMG 23695) which did not amplify. No amplicons were observed when other Vibrio species were tested (Table 1), including V. tubiashii ATCC 19109T. The amplicons obtained for the vcpA genes were verified by sequencing and confirmed the specificity of this assay for V. coralliilyticus strains. The same strain specificity was observed using the real-time PCR primer set (VcpARTF and VcpARTR) in standard PCR, with only one size of amplicon (166 bp) obtained for V. coralliilyticus isolates, with a mean melting temperature of 83.41°C±0.15. For products from isolates BH6, C1, C2 and TAV24, the melt curve analysis revealed a different peak corresponding to a melting temperature of 84.34°C±0.11. Using these same primers in reverse transcription real-time PCR with cDNA from V. coralliilyticus strains, CT values of 8–14 cycles were obtained, whilst other non-V. coralliilyticus species were detected at CT values of 21–31 cycles (Table 1). The RT-PCR result for V. coralliilyticus strain P3 (CT value of 25.67±0.58) confirmed the result observed in the standard PCR approach. The CT values for the no template control (NTC) were between 28–33 cycles.

The vcpA real-time PCR (VcpARTF-VcpARTR) specifically detected only V. coralliilyticus strains when a non-target culture of V. tubiashii ATCC 19109T was mixed in defined ratios (0∶100, 25∶75, 50∶50, 75∶25 and 100∶0%, respectively) after both target and non-target cultures were grown overnight and normalised to an OD600 of 1.0 (approximately 8×108 cells) and total DNA subsequently extracted and amplified (Fig. 1). The target V. coralliilyticus strains were detected at similar CT values (10.65–13.01) to those observed previously (Table 1) and CT values were positively correlated with V. coralliilyticus proportions (R2 = 0.956), indicating that non-target V. tubiashii did not interfere with vcpA gene amplification. Both 100% V. tubiashii and NTC CT values (25.96±1.02 and 30.45±1.20, respectively) were also within expected ranges.

Figure 1. Real-time PCR detection of V. coralliilyticus P1 cells in mixed cultures containing proportional numbers of V. tubiashii 19109T cells.

Each mixed culture was sampled in triplicate.

The specificity of the vcpA primers was validated by amplification of the same target and non-target genomic DNA mixtures using real-time PCR primers for the V. tubiashii zinc-metalloprotease vtpA [33]. These primers matched the V. coralliilyticus vcpA sequence in silico and so we intended to demonstrate that whilst the vcpA primers could detect changes in V. coralliilyticus concentration in the template mixture, the vtpA primers would amplify both the target and V. tubiashii non-target templates and so CT values would be similar for all mixtures. However, when the template mixtures were amplified by real-time PCR using the vtpA forward and reverse primers, we obtained the same result as for the vcpA primers; namely that CT values were positively correlated with V. coralliilyticus template DNA concentrations whilst 100% V. tubiashii ATCC 19109T and NTC CT values were again similar (data not shown).

Gharaibeh et al. (2009) developed the vtpA primers using the closely related V. tubiashii strains RE22 and ATCC 19105 [36]. Real-Time PCR of RE22 using both vtpA and vcpA primers resulted in CT values similar to those for V. coralliilyticus P1, at 10.96±0.16 and 8.91±0.05, respectively. Closer examination of nucleotide sequence data for all published metalloprotease genes for V. coralliilyticus (including sequences from two genomes, P1 and BAA-450) and V. tubiashii (with sequences from the genomes for strains ATCC 19109T and 19106) (Fig. 2) revealed significant differences between the V. tubiashii strains; as described previously [20], V. coralliilyticus has three copies of the metalloprotease (defined here as Types I, II and III) whereas V. tubiashii strains ATCC 19106 and 19109T have only one copy (Type I). V. tubiashii strain ATCC 19105 has one published Type I metalloprotease (VtpA) whilst RE22 however has two published metalloprotease gene copies (vtpA and vtpB) [37] grouped within Types I and II, respectively, and these are more closely related to metalloproteases from V. coralliilyticus strains than V. tubiashii strains ATCC 19109T and ATCC 19106 (Fig. 2). Recently, the unannotated de novo assembly of the genome of V. tubiashii RE22 has been published online Sequence comparisons of eight housekeeping gene loci (ftsZ, gapA, gyrB, mreB, pyrH, recA, rpoA and topA) from V. coralliilyticus BAA-450 and V. tubiashii strains ATCC 19106 and 19109T with the RE22 assembly revealed that the mean identity of matches with BAA-450 sequences was 98%, whilst with sequences from strains ATCC 19106 and 19109T, it was 88%. In a search of the RE22 assembly for the V. coralliilyticus BAA-450 Type III metalloprotease (Accession No. ZP_05885140.1), we also discovered a contig (RE22_contig_1616) containing a 1553 bp region with a 98% sequence identity to this gene.

Figure 2. Phylogenetic tree (using the neighbour-joining method) of Type I (1.8 kb), II (2.3 kb) and III (2 kb) metalloprotease genes from V. coralliilyticus and V. tubiashii (with V. splendidus included as an outgroup).

Figures at nodes indicate bootstrap values for 10000 bootstrap repetitions.


The Vibrionaceae are abundant in aquatic environments and are often found in high densities in association with marine organisms [38][40]. Historically, these opportunistic pathogens have long been associated with diseases of animals reared for aquaculture [41][45] but recently, the evidence for their increasingly important role in coral disease has been growing [2], [3] and a number of putative candidates have been studied, including V. shilonii [39], [46][50], V. harveyi [51], [52] and V. coralliilyticus [5], [8][21]. The relationship of the metalloproteases to pathogenicity amongst the prokaryotes is now well accepted [22] and their efficacy as virulence factors has been studied in a number of Vibrio species including V. tubiashii [37], [53], V. splendidus [54], [55], V. vulnificus [56][59] and V. anguillarum [60]. The zinc-metalloprotease of V. coralliilyticus has been shown to play an important role in the organism's virulence towards corals and their symbiotic zooxanthellae [8], [15], [20] and therefore this metalloprotease gene (vcpA) provided an attractive target for a detection assay. This study successfully developed specific primers to detect the vcpA gene in V. coralliilyticus using both PCR and reverse transcription real-time PCR approaches.

The PCR and real-time PCR primer sets designed in this study were able to specifically and routinely amplify DNA from all putative V. coralliilyticus isolates (Table 1), including a newly-described isolate (TAV24) infecting gorgonians in the Mediterranean Sea [61]. The specificity of the assays were such that they confirmed the misidentification of an isolate (P3) as V. coralliilyticus from a previous study [15] and also amplified an isolate (C2) which could not be detected by the real-time PCR assay of Pollock et al. (2010). Interestingly, melting curve analyses of the global V. coralliilyticus isolates revealed that the melting curve profiles for strains BH6, C1, C2 and TAV24 were different from all other V. coralliilyticus strains and this is correlated with the sequence divergence of their metalloproteases, as shown in a previous study [18]. When the GC content, which determines the melting curve profile, of the entire vcpA ORF nucleotide sequences derived from all global isolates was examined, a difference of >0.5% between strains BH6, C1, C2 and TAV24 (47.50±0.11%) and other V. coralliilyticus strains (46.90±0.16%) was observed. We are currently investigating the virulence of these two divergent clades of metalloproteases in V. coralliilyticus and melting curve profile analyses provide a rapid means for classification of these genes from newly sourced isolates.

A significant finding of the study was the serendipitous discovery of the shared metalloprotease and potential misclassification of V. tubiashii strains ATCC 19105 and RE22. The strain ATCC 19105 was first described by Tubiash et al. [62] as V. anguillarum and deposited in the American Type Culture Collection along with strains ATCC 19106 and ATCC 19109T. In a later study by Hada et al. [63], the three strains were reclassified as V. tubiashii using DNA-DNA hybridisation; interestingly, whilst ATCC 19106 shared a 96% DNA homology with ATCC 19109T, ATCC 19105 had only an 83% degree of reassociation with the type strain. Similarly, in comparative analyses of strains ATCC 19109T and ATCC 19105, there were notable differences in both the outer-membrane proteins produced and the expression of siderophores in iron utilisation [64], [65]. The historical confusion that surrounds identification of V. coralliilyticus and V. tubiashii species was first highlighted in the original article describing V. coralliilyticus [9], whereby the authors reclassified V. tubiashii LMG 10953 as V. coralliilyticus (strain BH5; Table 1). Following the incongruous quantitative PCR results obtained for V. tubiashii strains ATCC RE22 and 19109T, we constructed a phylogenetic tree using the metalloprotease sequence data for V. coralliilyticus and V. tubiashii strains (Fig. 2). V. coralliilyticus has three metalloproteases (Types I, II and III) [20], however, the recently published genomes for V. tubiashii strains ATCC 19106 [66] and 19109T (Accession Number NZ_AFWI00000000.1) identified each having only a single Type I metalloprotease. To date, the genome for ATCC 19105 has not been sequenced but two metalloproteases, vtpA [37] and vtpB [67] (Types I and II, respectively) have been characterised in the closely-related strain RE22 and these can be seen to cluster with those of V. coralliilyticus; the Type III metalloprotease has also been found within the unannotated RE22 genome recently published online The ATCC 19105 Type I and RE22 Type I and Type II metalloproteases shared >98% sequence identity with V. coralliilyticus strains ATCC BAA-450 and P1, compared with <72% for the Type I metalloproteases of V. tubiashii strains ATCC 19106 and 19109T. Whilst the presence of highly similar (>98%) multiple zinc-metalloprotease genes (and in the case of RE22, a number of other gene loci) strongly suggests a relatedness of ATCC 19105 and RE22 to V. coralliilyticus, further genomic sequence data is required to confirm their relationships and this is currently under investigation. Certainly, the virulence factors and mechanisms of pathogenesis studied independently in the shellfish pathogens RE22 and ATCC 19105 and coral pathogen V. coralliilyticus appear to be identical. Two isolates of V. coralliilyticus (BH5 and BH6) were previously obtained from diseased bivalve larvae (in the UK and Brazil, respectively) [9] but there is no evidence to confirm whether these strains were involved in shellfish pathogenesis. Since ATCC 19105 and RE22 have been shown to be major commercial pathogens of bivalves [37], if whole genome sequence data confirm their suspected identities, it would seem that the threat of V. coralliilyticus is no longer limited to coral reefs.

Due to the relative homogeneity of the 16S rRNA gene in Vibrio species, alternative gene targets for identification have been investigated and indeed the dnaJ gene has been suggested as a promising candidate [34], [68]. Pollock et al. (2010) previously designed a real-time PCR assay with TaqMan fluorescent probes for the dnaJ gene to detect V. coralliilyticus in environmental samples which was successful in specifically detecting twelve of thirteen tested V. coralliilyticus isolates. However, the assay developed during this current study improves upon this previous work due to its increased efficacy for discriminating V. coralliilyticus strains. The current assay was also optimised for analysis of mRNA, facilitating quantification of in situ gene expression in viable organisms, in contrast with genomic DNA which can be derived from extant or non-viable cells or even directly from seawater [69]. In their role of opportunist pathogens, Vibrio species are also known to comprise the normal €resident€ microbial communities associated with healthy corals [70] and so expression of a virulence factor such as the metalloprotease is useful in assessing the potential for a disease event. Indeed, zinc-metalloproteases have been targeted previously to monitor virulence during pathogenesis of oysters [33].

Conventional PCR has been used as a reliable molecular detection system for a variety of organisms in recent decades but is being replaced in many laboratories by real-time PCR analysis. Whilst the PCR method is time-consuming and methods of amplicon characterisation are laborious and not as sensitive as real-time PCR, equipment and consumable costs are much lower and therefore the technology remains more accessible to research laboratories. Additionally, conventional PCR allows amplification of much larger gene products than does real-time PCR and sequencing of products yields greater phylogenetic information. As we have shown previously, global V. coralliilyticus isolates contain one of two divergent Type I metalloprotease clades [18] and so classification of isolates requires that the entire vcpA open-reading frame be amplified by conventional PCR for meaningful sequence comparisons.

In conclusion, we successfully developed both PCR and reverse transcription real-time PCR assays for detection and quantification of the vcpA gene and mRNA, respectively, of V. coralliilyticus. While these assays do not require culturing and confirmation of individual isolates, they can obviously be used in concert with such techniques. The success of these assays in the presence of potentially interfering (and closely related) non-target DNA also suggests their use in ecological studies, where the target organism would likely be present amongst many other prokaryote and eukaryote cells. A significant result of this study is the discovery of the identical metalloproteases in V. tubiashii ATCC 19105 and RE22 strains causing disease in oysters and so these diagnostic tools may have important commercial applications in the aquaculture industry, as well as in reef management agencies, where early detection of virulent organisms would allow for the implementation of timely management steps to limit and possibly prevent losses due to V. coralliilyticus infection.

Materials and Methods

Bacterial strains, culture conditions and genomic DNA extraction

Global V. coralliilyticus isolates and strains of Vibrio species used in the study are listed in Table 1. Cultures of Vibrio species were grown overnight in Luria-Bertani broth plus 2% NaCl (LB20) on a horizontal shaker at 28°C and 200rpm. Total genomic DNA was extracted from overnight cultures using the Wizard Genomic DNA Kit (Promega, Madison, Wisconsin, USA) according to the manufacturers instructions.

PCR primer development

The vcpAF (5-ATG AAA CAA CGT CAA ATG CTT TG-3) and vcpAR (5-CCC TTT CAC TTC CGA TGT TGT G-3) PCR primers were previously designed to amplify the entire vcpA open reading frame (ORF) of V. coralliilyticus strain P1 and subsequently used to compare the vcpA genes from geographically disparate isolates of V. coralliilyticus [18]. The Lasergene EditSeq and MegAlign software packages (DNAStar, Madison, Wisconsin, USA) were used to check that the vcpAF and vcpAR primer pairs did not match any non-target Vibrio species. Primer sequences were also compared with those in the NCBI database using the Nucleotide Basic Local Alignment Search Tool (BLASTn) algorithm [35]. For quantitative real-time PCR primer design, the Lasergene Primer Select software package (DNAStar) was used to analyze potential primer pairs, and their specificity and range checked in silico as before. The chosen primer sequences for this real-time assay were vcpARTF (5-AGC TAC GAC TGC CGC CCT TAC-3) and vcpARTR (5-GGA GCC CTT TCA CTT ACG ATG TTG-3).

Construction of specific standard curves for Real-Time PCR

A standard curve for V. coralliilyticus quantification was generated by amplification of the vcpA ORF from V. coralliilyticus strain P1 genomic DNA using vcpAF forward and vcpARTR reverse primers. The amplicon was purified from agarose gels using the Nucleospin Extract II kit (Machery-Nagel, Dren, Germany), cloned into the pCR2.1-TOPO cloning vector and transformed into OneShot TOP10 competent cells as specified by the manufacturer (Invitrogen, Carlsbad, California, USA). Plasmids were prepared from transformants using the Qiagen Plasmid Purification kit (Qiagen, Valencia, California, USA) and insert size verified by agarose gel electrophoresis using SYBR Safe DNA gel stain (Invitrogen). The Lasergene SeqBuilder software package (DNAStar) was used to assess suitable restriction sites for linearisation of the plasmids such that digests did cut within the insert sequence. Plasmid DNA was quantified using a NanoDrop Spectrophotometer (Thermo Scientific, Wilmington, Delaware, USA) prior to digestion with Bgl II (New England Biolabs, Ipswich, Massachusetts, USA). Insert integrity was verified by PCR amplification using vcpAF and vcpARTR primers. The plasmids were serially diluted to prepare solutions containing 100–107 copies.

PCR amplification

Reaction mixtures comprised 1.25 mM MgCl2, 200 µM dNTP, 2 U of iTaq DNA Polymerase (Biorad, Hercules, USA), 200 nM of each primer, 200 ng µL−1 non-acetylated BSA [71], 1 µL (2–10 ng) of genomic DNA, and nuclease-free water to bring the total volume to 50 µL. For the vcpAF and vcpAR primer pair, reactions were initially denatured for 2 min at 94°C, followed by 30 cycles of denaturation at 94°C for 1 min, primer annealing at 64°C for 1 min and extension at 72°C for 2 min 30 s. This was followed by a final extension step of 72°C for 10 min. Amplicons were visualised by agarose gel electrophoresis using SYBR Safe DNA gel stain (Invitrogen).

RNA extraction, cDNA synthesis and real-time PCR amplification

Total RNA was extracted from early stationary phase cultures using the RiboPure Bacteria Kit (Ambion, Austin, Texas, USA). Reverse transcription was performed with the SuperScript III First Strand Synthesis System for RT-PCR (Invitrogen, Carlsbad, California, USA) as described by the manufacturer, using random hexamers to generate initial cDNA strands, which were quantified using a NanoDrop Spectrophotometer (Thermo Scientific). The real-time PCR assay comprised of 12.5 µL Qiagen Rotor-Gene SYBR Green RT-PCR Master Mix (Qiagen), 2.5 µL (10 µM) each of vcpARTF and vcpARTR primers and 2.5 µL nuclease-free water. Five microlitres of template (10 ng µL−1f either cDNA or plasmid DNA) was then added and each reaction performed in triplicate. A Qiagen Rotor-Gene Q real-time PCR cycler (Qiagen) was programmed with a modified Qiagen cycling program of a PCR initial activation step of 95°C for 5 min, followed by 40 cycles of denaturation of 95°C for 5 secs and a combined annealing and primer extension of 60°C for 30 secs. Quantitative PCR data was analysed using the Rotor-Gene Q software package (Qiagen).

Sequence Analysis

PCR products were purified from agarose gels using the Nucleospin Extract II kit (Machery-Nagel) and sent to Macrogen Inc. (Seoul, Korea) for sequencing. Sequences were trimmed and contigs assembled using the Lasergene SeqMan II software package (DNAStar). The neighbour-joining method (with the ClustalW package in MegAlign) was used to construct phylogenetic trees; V. splendidus was included as an outgroup. Gene accession numbers for metalloprotease sequence data used for phylogeny are listed in Table S1. Sequence data for complete vcpA genes for V. coralliilyticus strains were submitted under GenBank Accession Numbers JQ345033-JQ345046.


We would like to thank Frédérique Le Roux and David Goudenège (Station Biologique de Roscoff) for their assistance with comparative genomics analyses during the preparation of this manuscript, Jack Gilbert (University of Chicago) and Ben Temperton (Plymouth Marine Laboratory) for providing access to the Vibrio tubiashii ATCC 19106 genome sequence, Fabiano Thompson (Universidade Federal do Rio de Janeiro) for the Vibrio coralliilyticus P1 genome sequence and Pamela Morris (Belle W. Baruch Institute for Marine and Coastal Sciences) for providing strains. We would especially like to thank Claudia Häse (Oregon State University) for her assistance with the Vibrio tubiashii strain elucidation.

Author Contributions

Conceived and designed the experiments: BW DB. Performed the experiments: BW AM MB CHS. Analyzed the data: BW. Contributed reagents/materials/analysis tools: LV. Wrote the paper: BW DB.


  1. 1. Moberg F, Folke C (1999) Ecological goods and services of coral reef ecosystems. Ecol Econ 29: 215–233.
  2. 2. 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.
  3. 3. Bourne DG, Garren M, Work TM, Rosenberg E, Smith GW, et al. (2009) Microbial disease and the coral holobiont. Trends Microbiol 17: 554–562.
  4. 4. Lesser MP, Bythell JC, Gates RD, Johnstone R, Hoegh-Guldberg O (2007) Are infectious diseases really killing corals? Alternative interpretations of the experimental and ecological data. J Exp Mar Biol Ecol 346: 36–44.
  5. 5. Ben-Haim Y, Rosenberg E (2002) A novel Vibrio sp. pathogen of the coral Pocillopora damicornis. Mar Biol 141: 47–55.
  6. 6. Sussman M, Willis BL, Victor S, Bourne DG (2008) Coral pathogens identified for White Syndrome (WS) epizootics in the Indo-Pacific. PLoS ONE 3: e2393.
  7. 7. Arotsker L, Siboni N, Ben-Dov E, Kramarsky-Winter E, Loya Y, et al. (2009) Vibrio sp. as a potentially important member of the Black Band Disease (BBD) consortium in Favia sp. corals. FEMS Microbiol Ecol 70: 515–524.
  8. 8. Ben-Haim Y, Zicherman-Keren M, Rosenberg E (2003) Temperature-regulated bleaching and lysis of the coral Pocillopora damicornis by the novel pathogen Vibrio coralliilyticus. Appl Environ Microbiol 69: 4236–4242.
  9. 9. Ben-Haim Y (2003) Vibrio coralliilyticus sp. nov., a temperature-dependent pathogen of the coral Pocillopora damicornis. Int J Syst Evol Microbiol 53: 309–315.
  10. 10. Efrony R, Loya Y, Bacharach E, Rosenberg E (2006) Phage therapy of coral disease. Coral Reefs 26: 7–13.
  11. 11. Geffen Y, Ron EZ, Rosenberg E (2009) Regulation of release of antibacterials from stressed scleractinian corals. FEMS Microbiol Lett 295: 103–109.
  12. 12. Munn CB, Marchant HK, Moody AJ (2008) Defences against oxidative stress in Vibrios associated with corals. FEMS Microbiol Lett 281: 58–63.
  13. 13. Boroujerdi AFB, Vizcaino MI, Meyers A, Pollock EC, Huynh SL, et al. (2009) NMR-based microbial metabolomics and the temperature-dependent coral pathogen Vibrio coralliilyticus. Environ Sci Tech 43: 7658–7664.
  14. 14. Meron D, Efrony R, Johnson WR, Schaefer AL, Morris PJ, et al. (2009) Role of agella in virulence of the coral pathogen Vibrio coralliilyticus. Appl Environ Microbiol 75: 5704–5707.
  15. 15. Sussman M, Mieog JC, Doyle J, Victor S, Willis BL, et al. (2009) Vibrio zinc-metalloprotease causes photoinactivation of coral endosymbionts and coral tissue lesions. PLoS ONE 4: e4511.
  16. 16. Liu GF, Wang JY, Xu LW, Ding X, Zhou SN (2010) Sensitive and rapid detection of Vibrio corallilyticus by loop-mediated isothermal amplification targeted to the alpha subunit gene of RNA polymerase. Lett Appl Microbiol 51: 301–307.
  17. 17. Pollock FJ, Morris PJ, Willis BL, Bourne DG (2010) Detection and quantification of the coral pathogen Vibrio coralliilyticus by real-time PCR with TaqMan uorescent probes. Appl Environ Microbiol 76: 5282–5286.
  18. 18. Pollock FJ, Wilson B, Johnson WR, Morris PJ, Willis BL, et al. (2010) Phylogeny of the coral pathogen Vibrio coralliilyticus. Environ Microbiol Rep 2: 172–178.
  19. 19. Vizcaino MI, Johnson WR, Kimes NE, Williams K, Torralba M, et al. (2010) Antimicrobial resistance of the coral pathogen Vibrio coralliilyticus and Caribbean sister phylotypes isolated from a diseased octocoral. Microb Ecol 59: 646–657.
  20. 20. de O Santos E, Alves N, Dias G, Mazotto A, Vermelho A, et al.. (2011) Genomic and proteomic analyses of the coral pathogen Vibrio coralliilyticus reveal a diverse virulence repertoire. ISME J: 1–13.
  21. 21. Vidal-Dupiol J, Ladrière O, Meistertzheim AL, Fouré L, Adjeroud M, et al. (2011) Physiological responses of the scleractinian coral Pocillopora damicornis to bacterial stress from Vibrio coralliilyticus. J Exp Biol 214: 1533–1545.
  22. 22. Miyoshi S, Shinoda S (2000) Microbial metalloproteases and pathogenesis. Microbes Infect 2: 91–98.
  23. 23. Shao CP, Hor LI (2001) Regulation of metalloprotease gene expression in Vibrio vulnificus by a Vibrio harveyi LuxR homologue. J Bacteriol 183: 1369–1375.
  24. 24. Kawase T, Miyoshi S, Sultan Z, Shinoda S (2004) Regulation system for protease production in Vibrio vulnificus. FEMS Microbiol Lett 240: 55–59.
  25. 25. Miyoshi S (2006) Vibrio vulnificus infection and metalloprotease. J Dermatol 33: 589–595.
  26. 26. Ruwandeepika H, Bhowmick P, Karunasagar I, Bossier P, Defoirdt T (2011) Quorum sensing regulation of virulence gene expression in Vibrio harveyi in vitro and in vivo during infection of gnotobiotic brine shrimp larvae. Environ Microbiol Rep 3: 597–602.
  27. 27. Wang H, Wu J, Ayala J, Benitez J, Silva A (2011) Interplay among Cyclic Diguanylate, HapR, and the General Stress Response Regulator (RpoS) in the Regulation of Vibrio cholerae Hemagglutinin/Protease. J Bacteriol 193: 6529–6538.
  28. 28. Pollock F, Morris P, Willis B, Bourne D (2011) The Urgent Need for Robust Coral Disease Diagnostics. PLoS Pathog 7: e1002183.
  29. 29. Work TM, Richardson LL, Reynolds TL, Willis BL (2008) Biomedical and veterinary science can increase our understanding of coral disease. J Exp Mar Biol Ecol 362: 63–70.
  30. 30. Goarant C, Merien F (2006) Quantification of Vibrio penaeicida, the etiological agent of Syndrome 93 in New Caledonian shrimp, by real-time PCR using SYBR Green I chemistry. J Microbiol Meth 67: 27–35.
  31. 31. Blackstone GM, Nordstrom JL, Bowen MD, Meyer RF, Imbro P, et al. (2007) Use of a real time PCR assay for detection of the ctxA gene of Vibrio cholerae in an environmental survey of Mobile Bay. J Microbiol Meth 68: 254–259.
  32. 32. Gordon KV, Vickery MC, DePaola A, Staley C, Harwood VJ (2008) Real-time PCR assays for quantification and differentiation of Vibrio vulnificus strains in oysters and water. Appl Environ Microbiol 74: 1704–1709.
  33. 33. Gharaibeh DN, Hasegawa H, Häse CC (2009) Development of a quantitative real-time PCR assay for detection of Vibrio tubiashii targeting the metalloprotease gene. J Microbiol Meth 76: 262–268.
  34. 34. Saulnier D, De Decker S, Haffner P (2009) Real-time PCR assay for rapid detection and quantification of Vibrio aestuarianus in oyster and seawater: a useful tool for epidemiologic studies. J Microbiol Meth 77: 191–197.
  35. 35. Altschul S, Gish W, Miller W, Myers E, Lipman D (1990) Basic local alignment search tool. J Mol Biol 215: 403–410.
  36. 36. Estes RM, Friedman CS, Elston RA, Herwig RP (2004) Pathogenicity testing of shellfish hatchery bacterial isolates on Pacific oyster Crassostrea gigas larvae. Dis Aquat Organ 58: 223–230.
  37. 37. Hasegawa H, Lind EJ, Boin MA, Häse CC (2008) The extracellular metalloprotease of Vibrio tubiashii is a major virulence factor for pacific oyster (Crassostrea gigas) larvae. Appl Environ Microbiol 74: 4101–4110.
  38. 38. Nyholm SV, Stabb EV, Ruby EG, McFall-Ngai MJ (2000) Establishment of an animal-bacterial association: recruiting symbiotic Vibrios from the environment. Proc Natl Acad Sci USA 97: 10231–10235.
  39. 39. Sussman M, Loya Y, Fine M, Rosenberg E (2003) The marine fireworm Hermodice carunculata is a winter reservoir and spring-summer vector for the coral-bleaching pathogen Vibrio shiloi. Environ Microbiol 5: 250–255.
  40. 40. Thompson FL, Iida T, Swings J (2004) Biodiversity of Vibrios. Microbiol Mol Biol Rev 68: 403–431.
  41. 41. Sudheesh PS, Xu HS (2001) Pathogenicity of Vibrio parahaemolyticus in tiger prawn Penaeus monodon Fabricius: possible role of extracellular proteases. Aquaculture 196: 37–46.
  42. 42. Vandenberghe J, Thompson FL, Gomez-Gil B, Swings J (2003) Phenotypic diversity amongst Vibrio isolates from marine aquaculture systems. Aquaculture 219: 9–20.
  43. 43. Austin B, Austin D, Sutherland R, Thompson F, Swings J (2005) Pathogenicity of Vibrios to rainbow trout (Oncorhynchus mykiss, Walbaum) and Artemia nauplii. Environ Microbiol 7: 1488–1495.
  44. 44. Kesarcodi-Watson A, Kaspar H, Lategan MJ, Gibson L (2009) Two pathogens of Greenshell mussel larvae, Perna canaliculus: Vibrio splendidus and a V. coralliilyticus/neptunius-like isolate. J Fish Dis 32: 499–507.
  45. 45. Cano-Gómez A, Goulden EF, Owens L, Hφj L (2010) Vibrio owensii sp. nov., isolated from cultured crustaceans in Australia. FEMS Microbiol Lett 302: 175–181.
  46. 46. Banin E, Israely T, Kushmaro A, Loya Y, Orr E, et al. (2000) Penetration of the coral-bleaching bacterium Vibrio shiloi into Oculina patagonica. Appl Environ Microbiol 66: 3031–3036.
  47. 47. Banin E, Khare S, Naider F, Rosenberg E (2001) Proline-rich peptide from the coral pathogen Vibrio shiloi that inhibits photosynthesis of zooxanthellae. Appl Environ Microbiol 67: 1536–1541.
  48. 48. Kushmaro A, Banin E, Loya Y, Stackebrandt E, Rosenberg E (2001) Vibrio shiloi sp. nov., the causative agent of bleaching of the coral Oculina patagonica. Int J Syst Evol Microbiol 51: 1383–1388.
  49. 49. Banin E, Vassilakos D, Orr E, Martinez RJ, Rosenberg E (2003) Superoxide dismutase is a virulence factor produced by the coral bleaching pathogen Vibrio shiloi. Curr Microbiol 46: 418–422.
  50. 50. Nissimov J, Rosenberg E, Munn C (2009) Antimicrobial properties of resident coral mucus bacteria of Oculina patagonica. FEMS Microbiol Lett 292: 210–215.
  51. 51. Luna G, Biavasco F, Danovaro R (2007) Bacteria associated with the rapid tissue necrosis of stony corals. Environ Microbiol 9: 1851–1857.
  52. 52. Luna GM, Bongiorni L, Gili C, Biavasco F, Danovaro R (2009) Vibrio harveyi as a causative agent of the White Syndrome in tropical stony corals. Environ Micro Rep 2: 120–127.
  53. 53. Hasegawa H, Häse CC (2009) The extracellular metalloprotease of Vibrio tubiashii directly inhibits its extracellular haemolysin. Microbiol 155: 2296–2305.
  54. 54. Le Roux F, Binesse J, Saulnier D, Mazel D (2007) Construction of a Vibrio splendidus mutant lacking the metalloprotease gene vsm by use of a novel counterselectable suicide vector. Appl Environ Microbiol 73: 777–784.
  55. 55. Binesse J, Delsert C, Saulnier D, Champomier-Vergès MC, Zagorec M, et al. (2008) Metalloprotease vsm is the major determinant of toxicity for extracellular products of Vibrio splendidus. Appl Environ Microbiol 74: 7108–7117.
  56. 56. Jeong KC, Jeong HS, Rhee JH, Lee SE, Chung SS, et al. (2000) Construction and phenotypic evaluation of a Vibrio vulnificus vvpE mutant for elastolytic protease. Infect Immun 68: 5096–5106.
  57. 57. Valiente E, Lee CT, Hor LI, Fouz B, Amaro C (2008) Role of the metalloprotease Vvp and the virulence plasmid pR99 of Vibrio vulnificus serovar E in surface colonization and fish virulence. Environ Microbiol 10: 328–338.
  58. 58. Valiente E, Padrós F, Lamas J, Llorens A, Amaro C (2008) Microbial and histopathological study of the Vibriosis caused by Vibrio vulnificus serovar E in eels: the metalloprotease Vvp is not an essential lesional factor. Microb Pathog 45: 386–393.
  59. 59. Jones MK, Oliver JD (2009) Vibrio vulnificus: disease and pathogenesis. Infect Immun 77: 1723–1733.
  60. 60. Milton DL, Norqvist A, Wolf-Watz H (1992) Cloning of a metalloprotease gene involved in the virulence mechanism of Vibrio anguillarum. J Bacteriol 174: 7235–7244.
  61. 61. Vezzulli L, Previati M, Pruzzo C, Marchese A, Bourne DG, et al. (2010) Vibrio infections triggering mass mortality events in a warming Mediterranean Sea. Environ Microbiol 12: 2007–2019.
  62. 62. Tubiash HS, Chanley PE, Liefson E (1965) Bacillary necrosis, a disease of larval and juvenile bivalve molluscs. I. Etiology and epizootiology. J Bacteriol 90: 1036–1044.
  63. 63. Hada H, West P, Lee J, Stemmler J, Colwell R (1984) Vibrio tubiashii sp. nov., a pathogen of bivalve mollusks. Int J Syst Bacteriol 34: 1–4.
  64. 64. Beaubrun JG, Kothary M, Curtis S, Flores N, Eribo B, et al. (2008) Isolation and Characterization of Vibrio tubiashii Outer Membrane Proteins and Determination of a toxR Homolog. Appl Environ Microbiol 74: 907–911.
  65. 65. Beaubrun JG, Gopinath G, Kothary M, Franco A, Curtis S, et al. (2011) Inuence of iron-chelated growth conditions on outer membrane protein production and virulence of Vibrio tubiashii. Food Microbiol 28: 1409–1413.
  66. 66. Temperton B, Thomas S, Tait K, Parry H, Emery M, et al. (2011) Permanent draft genome sequence of Vibrio tubiashii strain NCIMB 1337 (ATCC19106). Standards in Genomic Sciences 4: 183–190.
  67. 67. Hasegawa H, Häse CC (2009) TetR-type transcriptional regulator VtpR functions as a global regulator in Vibrio tubiashii. Appl Environ Microbiol 75: 7602–7609.
  68. 68. Nhung PH, Shah MM, Ohkusu K, Noda M, Hata H, et al. (2007) The dnaJ gene as a novel phylogenetic marker for identification of Vibrio species. Syst Appl Microbiol 30: 309–315.
  69. 69. Nielsen K, Johnsen P, Bensasson D, Daffonchio D (2007) Release and persistence of extracellular DNA in the environment. Environ Biosaf Res 6: 37–53.
  70. 70. Wilson B, Aeby GS, Work TM, Bourne DG (2012) Bacterial communities associated with healthy and Acropora white syndrome-affected corals from American Samoa. FEMS Microbiol Ecol 80: 509–520.
  71. 71. Kreader CA (1996) Relief of amplification inhibition in PCR with bovine serum albumin or T4 gene 32 protein. Appl Environ Microbiol 62: 1102–1106.