Bacterial diversity of the digestive gland of ... - Wiley Online Library

Journal of Applied Microbiology ISSN 1364-5072

ORIGINAL ARTICLE

Bacterial diversity of the digestive gland of Sydney rock oysters, Saccostrea glomerata infected with the paramyxean parasite, Marteilia sydneyi T.J. Green1 and A.C. Barnes1,2 1 The University of Queensland, Centre for Marine Studies, Brisbane, Australia 2 The University of Queensland, School of Integrative Biology, Brisbane, Australia

Keywords 16S rDNA, bacterial community, Marteilia, oyster, Saccostrea. Correspondence Timothy J. Green, The University of Queensland, Centre for Marine Studies, Brisbane 4072, Australia. E-mail: [email protected]

2009 ⁄ 2123: received 10 December 2009, revised 10 January 2010 and accepted 12 January 2010 doi:10.1111/j.1365-2672.2010.04687.x

Abstract Aims: To determine whether the infestation by the protozoan paramyxean parasite, Marteilia sydneyi, changes the bacterial community of the digestive gland of Sydney rock oysters, Saccostrea glomerata. Methods and Results: Six 16S rDNA clone libraries were established from three M. sydneyi-infected and three un-infected oysters. Restriction enzyme analysis followed by sequencing representative clones revealed a total of 23 different operational taxonomic units (OTUs) in un-infected oysters, comprising the major phyla: Firmicutes, Proteobacteria, Cyanobacteria and Spirocheates, where the clone distribution was 44, 36, 7 and 5%, respectively. Close to half of the OTUs are not closely related to any other hitherto determined sequence. In contrast, S. glomerata infected by M. sydneyi had only one OTU present in the digestive gland. Phylogenetic analysis of the 16S rDNA sequence reveals that this dominant OTU, belonging to the a-Proteobacteria, is closely related to a Rickettsiales-like prokaryote (RLP). Conclusions: The microbiota of the digestive gland of Sydney rock oysters is changed by infection by M. sydneyi, becoming dominated by a RLP, and generally less diverse. The bacterial community of un-infected S. glomerata differs from previous studies in that we identified the dominant taxa as Firmicutes and a-Proteobacteria, rather than heterotrophic c-Proteobacteria. Significance and Impact of the Study: This is the first culture-independent study of the microbiota of the digestive glands of edible oysters to the species level. The commercial viability of the Sydney rock oyster industry in Australia is currently threatened by Queensland Unknown disease and the changes in the bacterial community of S. glomerata corresponding with infection by M. sydneyi sheds further light on the link between parasite infection and mortality in this economically damaging disease.

Introduction Oysters are filter feeders and can filter up to 109 bacterial cells per hour, many of which may opportunistically colonize the oyster with a detrimental effect during times of stress (Paillard et al. 2004; Pruzzo et al. 2005). Stressful conditions can be attributed to sub-optimal water quality, pollutants, farm management practices, harmful algae, phytoplankton quality and ⁄ or disease such as parasite infections (Garnier et al. 2007; Aladaileh et al. 2008).

There are numerous reports of environmental stresses leading to mass mortalities of farmed bivalves because of the proliferation of opportunistic pathogenic bacteria within oyster tissues (Paillard et al. 2004; Garnier et al. 2007), and both mortality and loads of the bacterium, Vibrio splendidus within oyster tissues have been positively correlated to stress hormones in juvenile Pacific oysters, Crassostrea gigas (Lacoste et al. 2001). Surprisingly, very few studies have investigated the role bacteria play in mortality of stressed aquaculture species because of infection

ª 2010 The Authors Journal compilation ª 2010 The Society for Applied Microbiology, Journal of Applied Microbiology 109 (2010) 613–622

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Bacterial diversity of Sydney rock oyster

by macro-parasites (Company et al. 1999; Bowman and Nowak 2004; Embar-Gopinath et al. 2005; Sitja`-Bobadilla et al. 2006), as investigations into causative agent of mass mortalities on aquaculture farms is often considered to be multi-factorial in origin (Company et al. 1999). For example, analysis of bacteria associated with amoebic gill disease of Atlantic salmon, Salmo salar, found species of bacteria only associated with the disease (Bowman and Nowak 2004; Embar-Gopinath et al. 2008), and the presence of these particular bacteria on salmon gills enhanced the ability of amoebae to infect gills and cause lesions (Embar-Gopinath et al. 2005). The Sydney rock oyster, Saccostrea glomerata, is a commercially important aquaculture species in Australia. Since the 1970s, mass mortalities of farmed S. glomerata have occurred because of Queensland Unknown disease (QX), which has been attributed to the protozoan parasite, Marteilia sydneyi (Perkins and Wolf 1976). The parasite is known to infect oysters via the gills or labial palps (mouth parts) and migrates through the oyster to the digestive gland over several weeks (Kleeman et al. 2002). Here, the parasite undergoes cell-within-cell proliferation and occupies all available spaces between the host’s epithelial cells of the digestive gland (Kleeman et al. 2002). Finally, the parasite undergoes sporulation and releases mature spores into the digestive tubule lumen (Perkins and Wolf 1976). The remainder of the parasite’s life cycle is currently unknown. Early field observations led researchers to suspect the involvement of other host species in the transmission of M. sydneyi (Lester 1986), as direct transmission of M. sydneyi from infected S. glomerata to un-infected S. glomerata by numerous methods were all unsuccessful (Lester 1986). The mechanism of oyster mortality because of infection with M. sydneyi is also unknown, but believed to be because of disruption of epithelial lining of the digestive gland causing the oyster to starve to death. However, there are no current scientific studies supporting this. Interestingly, studies have shown that S. glomerata can be infected with M. sydneyi without mortality. PCR studies have found M. sydneyi within the digestive gland of rock oysters in a number of estuaries in New South Wales, Australia, where mass mortality events because of QX disease do not occur (Adlard and Wesche 2005). For example, during the 2005 ⁄ 2006 summer, 27% of S. glomerata in Wallas Lakes, had sporulating M. sydneyi infections of the digestive gland, but no oyster mortality was reported by farmers (Jane Frances, NSW Department of Primary Industries, personal communications). The aim of the present study was to determine whether opportunistic pathogenic bacteria colonize the digestive gland of S. glomerata during QX disease episodes, thereby leading to oyster mortality because of secondary infection. 614

T.J. Green and A.C. Barnes

The diversity of bacteria present in the digestive glands of S. glomerata infected with M. sydneyi was compared to that of un-infected oysters by phylogenetic analyses of the bacterial 16S rRNA genes using PCR and cloning. The use of a culture-independent technique is a logical approach to understand what sort of pathogenic and nonpathogenic bacteria are associated with the digestive gland of S. glomerata and how this is influenced by the parasite, M. sydneyi, as previous studies have found that cultivatable bacteria represent 50%) in these oysters, which was diagnosed to be caused by QX disease (sporulating M. sydneyi infection within the digestive gland – Stage C, Kleeman et al. 2002) using histology by veterinarians at the Queensland Department of Primary Industries and Fisheries (Veterinarian and Chemical Residue Laboratory). No sign of mortality or disease agents were identified in oysters already been farmed on the lease. Un-infected S. glomerata used in this study were of the same age and size as M. sydneyi-infected S. glomerata, and were currently already being farmed on the same lease within the Pimpama R. Both infected and un-infected S. glomerata used in the current study originated as natural-caught spat from the Macleay R. and Moreton Bay, SE Queensland, respectively (O’Connor and Dove 2009). Previous studies have shown that M. sydneyi cannot be transmitted from infected oysters to un-infected S. glomerata (Lester 1986) allowing the direct comparison of bacterial communities of infected and un-infected individuals that had shared the same environment for 1 month. Un-infected and infected S. glomerata were collected and transported to



The University of Queensland by air-conditioned vehicle within an hour of collection. Upon immediate arrival at the laboratory, the external surfaces of the shells of each S. glomerata were scrubbed clean in a 4% chlorine solution, sprayed with 70% ethanol and shucked using a sterile oyster knife. Using a sterile Petri dish and scalpel blade each time, digestive gland samples were dissected and divided into two samples. The first sample was frozen at )80C for later DNA extraction. The second sample was used for tissue imprints on glass slides for QX disease diagnosis (Kleeman et al. 2002). Marteilia sydneyi diagnosis Saccostrea glomerata were confirmed to be either infected or un-infected with sporulating M. sydneyi by examining tissue imprints of the digestive gland stained with Hemacolor (Merck, Darmstadt, Germany) using a compound microscope following the method of Kleeman et al. (2002). Prior to 16S rDNA clone library construction, the presence or absence of M. sydneyi was confirmed using the PCR diagnostic method developed by Kleeman and Adlard (2000) using primers Leg1 (5¢-CGATCTGTGTAGTCGGATTCCGA) and Pro2 (5¢-TCAAGGGACATCCAACGGTC). PCR of 25 ll were carried out containing 50 ng of DNA template, 1· PCR buffer, 100 nmol l)1 of each primer, 2Æ5 mmol l)1 MgCl2, 4 · 100 lmol l)1 deoxynucleoside triphosphates and 0Æ5 U of Taq polymerase (recombinant; Invitrogen, Carlsberg, USA). PCR parameters consisted of an initial denaturation of 95C for 5 min, followed by 35 cycles of 95C for 30 s, 55C for 30 s, 72C for 30 s and a final elongation of 72C for 5 min. DNA purified from an oyster confirmed to be infected with M. sydneyi by cytology was performed as a positive control. PCR products were separated by electrophoresis on a 1Æ2% agarose ⁄ EtBr gel alongside a low molecular weight DNA marker (Fermentas, Ontario, Canada). DNA extraction Community nucleic acids were extracted from the digestive glands of S. glomerata using a bead-beating and freeze– thaw protocol. For each oyster, a 1 mm3 sample of digestive gland tissue was transferred to a 2 ml screw cap tube containing 0Æ5 ml of lysis buffer (50 mmol l)1 Tris Base, 0Æ75 mol l)1 Sucrose, 40 mmol l)1 EDTA, pH 8Æ3) and acid washed glass beads (150–212 and 2500 lm beads). Digestive gland tissue was homogenized using a MagNA Lyser (Roche Diagnostics, Basel, Switzerland) at 3000 rev min)1 for 1 min. Lysozyme (to 5 mg ml)1) and RNase A (to 50 lg ml)1) were then added to each tube and incubated at 37C for 1 h. Samples were then subjected to three


rounds of freeze–thaw lysis before the addition of Proteinase K (to 1 mg ml)1) and sodium dodecyl sulfate [to 0Æ3% (w ⁄ v)] and incubated at 70C for 10 min. Samples were again subjected to three rounds of freeze–thaw lysis before being vortexed in the presence of one volume of phenol. Lysates were extracted with phenol–chloroform, then sodium acetate was added to 0Æ3 mol l)1, and nucleic acids were precipitated from solution by addition of one volume of isopropanol. Extracted DNA was purified by electrophoresis through a 1Æ2% low-melting agarose gel (Bioline, NSW, Australia) and DNA bands (‡10 kb in size) were excised from the gel and purified from the gel slice with a MegaSpin agarose gel extraction kit (iNtRON biotechnology, Korea) and eluted in 20 ll of 10 mmol l)1 Tris, pH 8Æ0. 16S rDNA amplification and cloning Six clone libraries were prepared from three QX-infected and three un-infected S. glomerata. Community ribosomal DNAs (rDNAs) were PCR amplified from 1 to 50 ng of bulk DNA in three parallel PCR per oyster digestive gland sample containing (as final concentrations) 1· high fidelity PCR buffer, 2Æ5 mmol l)1 MgCl2, 4 · 200 lmol l)1 deoxynucleoside triphosphates, 1 U of High Fidelity Platinum Taq DNA polymerase (Invitrogen) and 300 nmol l)1 each of the universal bacterial primers 27F (5¢-AGAGTTTGATCCTGGCTCAG) and 1492R (5¢-GTTACCTTGTTACGACTT). Dimethyl sulfoxide [DMSO, 5% (v ⁄ v) final concentration] was added to promote amplification of templates. Reaction mixtures were incubated in an Eppendorf Mastercycler (Eppendorf, Hamburg, Germany) with an initial denaturation of 94C for 5 min, followed by 27 cycles of 94C for 15 s, 49Æ5C for 1 min, 72C for 2 min with a final elongation of 72C for 5 min. Amplified rDNAs were pooled from three individual reactions per sample and purified (PCRquickspin PCR product purification kit; iNtRON biotechnology) and ligated overnight into pCR4 TOPO vector and transformed using TOP10 One Shot chemically competent cells (Invitrogen). Transformed cells were cultured on Luria–Bertani agar plates containing 100 mg l)1 of ampicillin and 120 clones from each library were randomly picked and inserts amplified by PCR using the Universal M13 primer set. Insert size was checked by agarose (1Æ2%) gel electrophoresis against a 1 kb DNA ladder (Fermentas). Restriction fragment length polymorphism (RFLP) screening of rDNA clones rDNA inserts from recombinant clones were double digested overnight at 37C with 0Æ1 U each of the


615



four-base-specific restriction endonucleases HinPI and MspI in 1· NEB Buffer 2 (New England Biolabs, Ipswich, MA). Digested products were separated by agarose (2Æ5%) gel electrophoresis. Bands were visualized by staining with ethidium bromide and UV illumination. RFLP patterns for each library were grouped visually, and up to three representatives from each RFLP pattern type were selected (Results) for sequencing. Sequencing of PCR products from cloned inserts rDNA inserts from representative clones (Table 1) were reamplified by PCR as above and residual primers and enzyme was removed using exonuclease I and shrimp alkaline phosphatase (Fermentas). PCR products were forward and reverse sequenced using the universal M13F and M13R primers by the Australian Genome Research Facility. Obtained sequences were quality scored and visually checked for possible sequencing errors and vector

sequences were removed using VecScreen (http:// www.ncbi.nlm.nih.gov/VecScreen). Phylogenetic analyses Sequences were initially compared to the available databases by using the blast (basic local alignment search tool) network service (Altschul et al. 1997) to determine their approximate phylogenetic affiliations and orientation. Partial sequences were then compiled in Vector NTI ver. 10 (Invitrogen) and chimeric sequences were removed using the Chimera Check analysis function of the Ribosomal Database Project (Cole et al. 2003). Sequences were then aligned with selected reference 16S rRNA sequences from GenBank using the ClustalW algorithm in the computer software program Mega ver. 4.0 (Tamura et al. 2007) and phylogenetic trees were constructed using the neighbour-joining distance method (Saitou and Nei 1987). The statistical significance of

Table 1 Summary of the 16S rDNA sequences identified in Saccostrea glomerata digestive glands from two clone libraries, QX-infected and un-infected

Type sequence QX library QX Clone 28 Un-infected library Control clone 2 Control Control Control Control Control Control Control Control Control

clone clone clone clone clone clone clone clone clone

82 22 87 26 69 67 108 12 64

Control Control Control Control Control Control Control Control Control Control Control Control Control

clone clone clone clone clone clone clone clone clone clone clone clone clone

16 9 48 93 120 23 92 98 118 10 47 83 109

Other sequence representatives

No. of clones

Bacteria division

Q16, Q17, Q26, Q27, Q45, Q46, Q102, Q103, Q104, Q116

120

a-Proteobacteria

C3, C4, C7, C8, C25, C34, C36, C42, C59, C66, C88, C95, C99 None None C41, C56 None None None None C21, C71 C4, C13, C17, C19, C20, C30, C35, C39, C63, C103, C116 C117 None None None C5 C68 None None C46 None None None None

26

a-Proteobacteria

2 1 4 1 1 1 1 3 37

a-Proteobacteria c-Proteobacteria c-Proteobacteria c-Proteobacteria Fusobacteria Firmicute Firmicute Firmicute Firmicute

2 1 1 1 2 1 1 1 3 1 1 2 1

Spirochaetes Spirochaetes Spirochaetes Spirochaetes Chlorophyta Cyanobacteria Cyanobacteria Cyanobacteria Cyanobacteria Cyanobacteria Actinobacteria Actinobacteria Chloroflexi

Database matches (>97% identity)

Accession no.

Sphinyomonsa sp.

AY081166

Vibrio sp. Cetobacterium somerae Dolsigranulum pigrum Clostridium ruminantium

AM159569 AB353124 NR_026098 EU089964

Uncultured Cyanobacterium Uncultured Acaryochloris sp Synechococcus sp. Synechococcus sp. Synechococcus sp. Propionibacterium acnes Propionibacterium acnes

DQ446127 DQ917811 AF448063 AY172826 CP000110 AB108481 AB042290

QX, Queensland Unknown.

616



interior nodes was determined by performing bootstrap analyses based on 1000 resamplings of the data. Nucleotide sequence accession numbers The 16S rDNA sequences from the infected and un-infected clone libraries were deposited in the EMBL database (Genbank accession nos. FM995169–FM995192). Sequences originating from the un-infected clone library are designated SRODG followed by the clone number, whereas, sequences from the infected clone library are designated SROQX followed by the clone number. Results QX disease confirmation Saccostrea glomerata used for library construction were confirmed to be either infected or un-infected by QX disease (M. sydneyi sporulating in the digestive gland). Un-infected S. glomerata were shown to be free of M. sydneyi within the digestive gland by PCR, whereas, PCR and tissue imprints confirmed the presence of several life stages of M. sydneyi including sporulating parasite within the digestive gland of infected S. glomerata (Fig. 1). Community analysis To determine whether the bacterial community of the digestive gland of S. glomerata is influenced by M. sydneyi infection, we analysed the 16S rRNA genes obtained by PCR from DNA purified from the digestive

Figure 1 Tissue imprints of the digestive gland of Saccostrea glomerata infected with Marteilia sydneyi showing mature sporonts (arrows). Scale bar, 10 lm. Hemacolor stain.


gland of M. sydneyi-infected and un-infected S. glomerata. As different DNA extraction procedures and PCR conditions can result in differential recovery of rRNA genes, we trialled several different methods for DNA extraction but found all other methods unsatisfactory with either poor yields, presence of inhibitory compounds or sheared DNA. Likewise, all attempts to amplify the 16S rDNA gene without DMSO were unsuccessful. To minimize PCR biasing, the number of PCR cycles were optimized to 27 cycles and three identical PCR for each sample were run at the same time to increase yield. To determine individual variation in the bacterial community between oysters, amplified rDNA from three M. sydneyi-infected and three uninfected oysters were ligated to generate six clone libraries. From these six libraries, a total of 720 clones containing inserts of the expected size (c. 1Æ5 kb) were selected for further analysis. Typically, 2–7 bands resulted from each rDNA restriction digest in the discernible fragment size range of 50– 500 bp (data not shown). For M. sydneyi-infected S. glomerata clone libraries, 3–5 different RFLP types were distinguished visually from each of the three libraries. For the un-infected S. glomerata clone libraries, 36–54 different RFLP types were distinguished visually from each of the three libraries. From the RFLP results, it was decided that RFLP representative from only one QX-infected and one un-infected S. glomerata bacterial clone library would need to be sequenced as the three QX-infected and the three un-infected bacterial clone libraries were representative of each other, respectively. For the infected and un-infected libraries, a total of 11 and 66 clones were sequenced representing the unique RFLP types determined for each library. The 16S rDNA sequences analysed, the frequencies with which they were obtained in the libraries, and their phylogenetic positions based on their 16S rDNA sequences are summarized in Table 1. A total of 23 different species from at least eight bacterial divisions were identified from the un-infected S. glomerata clone library (Fig. 2). Only half of the 16S rDNA gene sequences represented by 12 operational taxonomic units (OTUs) were affiliated with already existing sequences in the public databases with ‡97% identity. The remaining 11 sequences were remotely related to 16S rRNA sequences of known bacteria, and represent novel phylotypes not described in previous analyses. The majority of OTUs from the un-infected S. glomerata clone library are associated phylogenetically within the bacterial divisions of Firmicutes (44%), Proteobacteria (36%), Cyanobacteria (7%) and Spirochaetes (5%) of OTUs in the library. The remaining bacteria groups such as Actinobacteria, Chloroflexi, Chlorophyta and Fusobacteria make up a total of 8% of the OTUs within the library. In un-infected S. glomerata digestive


617



Anaplasma marginale AF414878 100 Anaplasma centrale AF414869 100 Anaplasma ovis AF414870 Anaplasma platys AF303467 Ehrlichia bovis EBU03775

100 95

50

Cowdria ruminatium X61768 Candidatus Xenohaliotis californiensis AF133090 Ehrlichia sennetsu M73225

56 92

100 Control clone 2 QX clone 28

80

a-Proteobacteria

Rickettsia rickettsii M21293 Thalassospira profundimaris AY186195 Control clone 90

99 74

100 50

99 98

Sphingomonadacae bacterium DQ490375 Uncultured Eubacterium AJ292591 Sphingomonas sp. AY081166 99 Control clone 82 Piscirickettsia salmonis U36941 Coxiella burnetii M21291

99

Shewanella alga AF005250

100

Control clone 22

100

Shewanella fidelia AF420312 Shewanella waksmanii AY170366

93 54

100

Vibrio sp. V125 DQ146977 Photobacterium phosphoreum AB179540

100 100

93

100 97

g -Proteobacteria

Control clone 87 Vibrio aestuarianus AJ845014 Vibrio vulnificus X76334 Vibrio sp. AM159569 Control clone 26 Vibrio ichthyoenteri AJ437192 Vibrio scophthalmi U46579 Cetobacterium ceti X78419 Control clone 69

100

Fusobacteria

98 Cetobacterium somerae AJ438155 100 Dolosigranulum pigrum X70907 Control clone 67

99

Bacillus sp. AY853168 Clostridium bifementans DQ978211

56

Control clone 108

100

74

Control clone 12

Firmicute

Control clone 64 Mycoplasma hyorhinis EU643797

100

99

99

Mycoplasma gypis AF125589 Mycoplasma spumans AF538684

93 100

84 85 98

Borrelia parkeri AF210137 Borrelia duttonii AF107366 Control clone 16

Spirochaeta aurantia AY599019 Spirochaeta halophila M88722

92 98

Spirochaete

Control clone 9 Uncultured Spirochaete sp. AY712343

70

Control clone 48 100 100

Chlorella sp. EF114678 Chlorella vulgaris X16579

Chlorophyta

Control clone 120 100 Uncultured cyanobacterium DQ446127 Control clone 23

54 100 99 93

73

Acaryochloris sp. DQ917811

100

Control clone 92 Acaryochloris marina AY163573 89 Synechococcus sp. AY183115 88

Control clone 98

Cyanobacteria

Uncultured cyanobacterium AM259252 Cyanobium sp. AY183115 Synechococcus sp. AY172826

100

Control clone 118 96

Uncultured cyanobacterium AM259793

94 Uncultured Synechoccus sp. AY125377 99 Control clone 10 100 Control clone 47 Propionibacterium acnes AB042290 79 Control clone 83

Actinobacteria

Chloroflexi Bacterium DQ351765 100

Control clone 109

Aquifex pyrophilus M83548 0·02

618

Chloroflexi Aquificae

Figure 2 Evolutionary distance dendrogram of the bacterial 16S rDNA sequence types obtained from Saccostrea glomerata digestive glands. Reference sequences were chosen to represent the broadest diversity of bacteria. Aquifex pyrophilus was used as outgroup for the analysis. Division level groupings are bracketed at the right o the figure.



glands, two dominant phylotypes were detected. The first grouped in the Firmicute division (42% of OTUs), while the other belonged to the a-Proteobacteria division (27% of OTUs). These two phylotypes constitute novel subclasses within the Firmicute and a-Proteobacteria divisions. The closest match in the available databases for the Firmicute phylotype was to Mycoplasma hyorhinis (86% identity). The a-Proteobacteria phylotype was similar to that of species found in the bacterial division of Rickettsiales, with similarity to Anaplasma marginale, Wolbachia pipentis, Rickettsia rickettsii and Candidatus Xenohaliotis californiensis, where the sequence identity was 87, 84, 83 and 83%, respectively. The 16S rDNA sequences for both the Firmicute and a-Proteobacteria phylotypes comprised gene clusters (multiple RFLP patterns and 16S rDNA sequences identified). Phylogenetic clusters of closely related but distinct 16S rRNA sequences (‡97% identity) are often observed in phylogenetic studies in marine samples (Field et al. 1997). To simplify the presentation of the phylogenetic tree (Fig. 2), gene clusters with ‡97% identity were reduced to one representative sequence (Table 1). However, gene clusters are thought to be biologically relevant, the result of sequence variability among ribosomal operons within a single strain or sets of closely related cellular lineages of organisms (Field et al. 1997); however, artefacts of PCR (polymerase error or chimera formation) cannot be ruled out. In contrast to un-infected oysters, all of the OTUs from the QX-infected S. glomerata clone library belonged to the novel subclass of a-Proteobacteria (‡99% identity, 100% of OTUs). Discussion There has been extensive research into the bacterial communities of edible oysters using culture-dependent techniques as opportunistic bacteria are known to cause mass mortalities of farmed bivalves during times of environmental stress (Paillard et al. 2004; Schulze et al. 2006; Garnier et al. 2007). Additionally, the risk of uncooked shellfish causing gastroenteritis in humans is well established (Kueh and Chan 1985; Pujalte et al. 1999; Richards et al. 2008). Relatively few bacteria in oysters are culturable (Romero et al. 2002), and an alternative to reliance on laboratory culture is the use of molecular approaches to identify microbial communities based on the phylogenetic analyses of their rRNA genes (Pace et al. 1985). There are relatively few reports of the use of culture-independent methods to describe bacterial diversity of apparently healthy (Romero and Espejo 2001; Romero et al. 2002; Hernańdez-Za´rate and Olmos-Soto 2006) or diseased bivalves (Boardman et al. 2008).


The bacterial diversity of the digestive gland is different in oysters infected by Marteilia sydneyi Results obtained in this study suggest that un-infected S. glomerata have a large bacterial diversity within their digestive gland with 23 different phylotypes identified from eight different bacterial divisions (Table 1). The majority of bacterial taxa identified in un-infected S. glomerata digestive glands were either documented marine bacteria and ⁄ or have been previously cultured from oyster tissues. Two dominant bacterial taxa were identified from the clone library constructed from un-infected S. glomerata digestive glands. Based on the phylogenetic analysis of the 16S rDNA sequences, the Firmicute and a-Proteobacteria are likely to be intracellular bacteria. Intracellular bacteria are common endosymbionts in molluscs, often un-associated with disease (Elston 1986). This finding is in contrast with other studies identifying c-Proteobacteria being the most frequently detected bacterial division in bivalves (Kueh and Chan 1985; Pujalte et al. 1999; Beleneva et al. 2003, 2007). The c-Proteobacteria group includes bacteria within Vibrionaceae, with Vibrio as the principle representative of the family. Vibrio species are the most commonly identified bacteria within shellfish, (Kueh and Chan 1985; Pujalte et al. 1999; Beleneva et al. 2007), as most studies into the bacterial communities of bivalves are biased towards identifying culturable heterotrophic bacteria. In the current study, three different species contained within the Vibrionaceae were identified from six clones representing a total of 6% of the OTUs. The low numbers of Vibrionaceae identified is not surprising in the current study as previous studies have shown that the most abundant bacteria in oysters are nonculturable (Romero et al. 2002) and bacteria in the digestive tract of oysters differ considerably from those isolated from the whole animal (Kueh and Chan 1985). A previous study using culture-independent techniques identified c-Proteobacteria and Grampositives with a low G+C content (includes Firmicutes) as the dominant metabolically active bacteria within the digestive gland of healthy C. gigas (Hernańdez-Za´rate and Olmos-Soto 2006). In contrast to healthy oysters, the digestive glands of M. sydneyi-infected S. glomerata were dominated by one phylotype. Based on the phylogenetic analysis of the 16S rDNA sequences, this phylotype is an a-Proteobacterium and shows high homology to other 16S rRNA sequences from Rickettsiale-like prokaryotes (RLP). This bacterium was also found to be one of the dominant species in un-infected oysters, but only represented a quarter of the clones in those libraries. RLPs are known to cause disease in oysters (Renault and Cochennec 1994; Wu and Pan 2000; Sun and Wu 2004). The sequence showed similarity


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to Candidatus X. californiensis (83% identity), the causative agent of withering syndrome of farmed and wild abalone on the West Coast of North America and in Europe (Friedman et al. 2000; Moore et al. 2001; Balseiro et al. 2006). Studies have also shown that Candidatus X. californiensis infects the gastrointestinal epithelia of the digestive gland in abalone. The disease is characterized by the degeneration of the digestive gland and cessation of feeding by the host, which slowly starves to death (Friedman et al. 2000). The symptoms of withering syndrome in abalone is similar to those attributed to QX disease in S. glomerata. Interestingly, there has been a report of the co-infection of the Haplosporidian parasite, Haplosporidium montforti and Candidatus X. californiensis resulting in the mass mortalities of the European abalone, Haliotis tuberculata (Balseiro et al. 2006). Further studies are required to determine whether the RLP found in digestive glands of QX-infected S. glomerata is not merely coincidental in the current experiment described here. This will require specific diagnostic tools such as PCR and in-situ hybridization to be developed to quantify the number of RLP that colonize the digestive gland of M. sydneyiinfected and un-infected oysters, and, does the presence of this novel RLP correlate with estuaries where mortality of S. glomerata occurs because of QX disease? Alternatively, the role of this RLP in the mortality of S. glomerata infected with M. sydneyi could be investigated by treating S. glomerata with oxytetracycline before exposing them to M. sydneyi, as the RLP is likely to be sensitive to oxytetracycline (Friedman et al. 2003). This study is supportive of the contention that S. glomerata infected by M. sydneyi cease feeding. Clone libraries constructed from un-infected S. glomerata revealed the presence of bacteria within the digestive gland associated with filter feeding (Cyanobacteria sp.) or associated with the natural micro flora of the digestive gland that are involved in digestion (Spirochaetes and Clostridium sp.). The absence of these bacteria within the digestive glands of QX-infected S. glomerata, suggests that the oysters have ceased feeding resulting in the bacterial community to be dominated by intracellular bacteria, allowing the oysters to starve to death. However, M. sydneyi infection did change the bacterial community because other intracellular bacteria (Firmicutes, Mycoplasma sp.) present in the un-infected clone library were absent in the M. sydneyiinfected clone library (Fig. 1). Conclusion QX disease is associated with changes in the bacterial community within the digestive gland of S. glomerata. Un-infected S. glomerata have a diverse range of bacteria within their digestive gland, but S. glomerata infected 620


with M. sydneyi results in the presence of one dominant bacterial type, a RLP. The effect of this bacterium on the host is currently unknown, but based the similarity of its 16S rDNA sequence to other known pathogens it is likely to have possible detrimental effects. Whether changes in the bacterial community of the digestive gland of S. glomerata is a result of infection by M. sydneyi or changes in the bacterial community allows infection by M. sydneyi is currently unknown and warrants further investigation. Acknowledgements The authors would like to acknowledge the assistance of H. Gobert and A. Chai. We acknowledge the Australian government for Australian Postgraduate Award to T. Green. References Adlard, R.D. and Wesche, S.J. (2005) Aquatic Animal Health Subprogram: Development of a Disease Zoning Policy for Marteilia sydneyi to Support Sustainable Production, Health Certification and Trade in the Sydney Rock Oyster. Final project report FRDC2001 ⁄ 214, pp. 1–46. Queensland Museum, Brisbane. Aladaileh, S., Nair, S.V. and Raftos, D.A. (2008) Effects of noradrenaline on immunological activity in Sydney rock oysters. Dev Comp Immunol 32, 627–636. Altschul, S.F., Madden, T.L., Scha¨ffer, A.A., Zhang, J., Zhang, Z., Miller, W. and Lipman, D.J. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 17, 3389–3402. Balseiro, P., Aranguren, R., Gestal, C., Novoa, B. and Figueras, A. (2006) Candidatus Xenohaliotis californiensis and Haplosporidium montforti associated with mortalities of abalone Haliotis tuberculata cultured in Europe. Aquaculture 258, 63–72. Beleneva, I.A., Zhukova, N.V. and Maslennikova, E.F. (2003) Comparative study of microbial communities from cultured and natural populations of the mussel Mytilus trossulus in Peter the Great Bay. Microbiology 72, 472– 477. Beleneva, I.A., Zhukova, N.V., Lan, H.L. and Nguyen Tran, D.H. (2007) Taxonomic composition of bacteria associated with cultivated molluscs Crassostrea lugubris and Perna viridis and with the water of the Gulf of Nha Trang Lagoon, Vietnam. Microbiology 76, 220–228. Boardman, C.L., Maloy, A.P. and Boettcher, K.J. (2008) Localisation of the bacterial agent of juvenile oyster disease (Roseovarius crassostreae) within affected Eastern oyster (Crassostrea virginica). J Invertebr Pathol 97, 150–158. Bowman, J.P. and Nowak, B. (2004) Salmonid gill bacteria and their relationship to amoebic gill disease. J Fish Dis 27, 483–492.



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