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Multilocus virulence typing of clinical and environmental Vibrio vulnificus isolates

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Title:
Multilocus virulence typing of clinical and environmental Vibrio vulnificus isolates
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Book
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English
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Gordon, Katrina V
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University of South Florida
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Real-time PCR
BOX PCR
ViuB
Vcg
16S rRNA
Dissertations, Academic -- Biology -- Doctoral -- USF   ( lcsh )
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non-fiction   ( marcgt )

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Summary:
ABSTRACT: The bacterium Vibrio vulnificus is an autochthonous inhabitant of estuarine waters and also found in shellfish such as oysters. It is a human pathogen of importance in the seafood industry, and can also infect recreational water users. Currently, recognized methods of detection rely upon isolation of pure cultures which requires at least 24 hours. To reduce the time needed for identification of the pathogen and simultaneously ascertain the virulence potential of the strains present, real-time PCR assays and sample processing procedures were developed (Chapter 1). These assays discriminate between type A (environmental, generally lower virulence) and type B (clinical, higher virulence) isolates.The genetic relationships between environmental V. vulnificus strains isolated from permitted and prohibited shellfish harvesting areas was determined using BOX-PCR genomic fingerprinting coupled with sequence analysis of three proposed virulence markers: (1) the virulence correlated gene (vcg), (2) 16S rRNA type and (3) presence/absence of the vulnibactin gene (viuB) (Chapter 2). The real-time PCR assays were able to detect the presence of seeded V. vulnificus in environmental water at a concentration of 160 cells 100·ml⁻¹. In seeded oyster homogenates, the assays were able to detect a minimum of 10³ cells and 10² cells per reaction of type A and type B respectively. The phylogenetic analysis separated the majority of type A/ vcgE strains isolated from permitted shellfish harvesting areas from those isolated from prohibited harvesting areas. The genomic (BOX-PCR) fingerprints of type A and type AB isolates were more similar to one another than to type B isolates.Only one type A/ vcgE isolate contained the viuB gene; however, eight type B/ vcgC isolates had that gene. No obvious grouping was discerned between type B/ vcgC isolates from permitted versus prohibited shellfish harvesting areas or between those possessing the viuB gene versus those lacking viuB. These data provide insight into the ecology and correlation between population biology and general water quality, as gauged by the classification of the shellfish growing area waters. The 16S typing assays can be used for routine rapid typing to aid in risk assessment and reduce infection frequency through consumption of contaminated seafood.
Thesis:
Dissertation (Ph.D.)--University of South Florida, 2008.
Bibliography:
Includes bibliographical references.
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Mode of access: World Wide Web.
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by Katrina V. Gordon.
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Title from PDF of title page.
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Document formatted into pages; contains 92 pages.
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Includes vita.

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aleph - 002021453
oclc - 428448953
usfldc doi - E14-SFE0002557
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ABSTRACT: The bacterium Vibrio vulnificus is an autochthonous inhabitant of estuarine waters and also found in shellfish such as oysters. It is a human pathogen of importance in the seafood industry, and can also infect recreational water users. Currently, recognized methods of detection rely upon isolation of pure cultures which requires at least 24 hours. To reduce the time needed for identification of the pathogen and simultaneously ascertain the virulence potential of the strains present, real-time PCR assays and sample processing procedures were developed (Chapter 1). These assays discriminate between type A (environmental, generally lower virulence) and type B (clinical, higher virulence) isolates.The genetic relationships between environmental V. vulnificus strains isolated from permitted and prohibited shellfish harvesting areas was determined using BOX-PCR genomic fingerprinting coupled with sequence analysis of three proposed virulence markers: (1) the virulence correlated gene (vcg), (2) 16S rRNA type and (3) presence/absence of the vulnibactin gene (viuB) (Chapter 2). The real-time PCR assays were able to detect the presence of seeded V. vulnificus in environmental water at a concentration of 160 cells 100ml. In seeded oyster homogenates, the assays were able to detect a minimum of 10 cells and 10 cells per reaction of type A and type B respectively. The phylogenetic analysis separated the majority of type A/ vcgE strains isolated from permitted shellfish harvesting areas from those isolated from prohibited harvesting areas. The genomic (BOX-PCR) fingerprints of type A and type AB isolates were more similar to one another than to type B isolates.Only one type A/ vcgE isolate contained the viuB gene; however, eight type B/ vcgC isolates had that gene. No obvious grouping was discerned between type B/ vcgC isolates from permitted versus prohibited shellfish harvesting areas or between those possessing the viuB gene versus those lacking viuB. These data provide insight into the ecology and correlation between population biology and general water quality, as gauged by the classification of the shellfish growing area waters. The 16S typing assays can be used for routine rapid typing to aid in risk assessment and reduce infection frequency through consumption of contaminated seafood.
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Multilocus Virulence Typing of Clinical and Environmental Vibrio vulnificus Isolates by Katrina V. Gordon A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy Department of Biology College of Arts and Sciences University of South Florida Major Professor: Valerie J. Harwood, Ph.D. Angelo DePaola Jr, Ph.D. James D. Garey, Ph.D. Daniel V. Lim, Ph.D. Date of Approval: July 18th 2008 Keywords: Real-time PCR, BOX PCR, viuB vcg 16S rRNA, genotyping Copyright 2008, Katrina V. Gordon

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ii ACKNOWLEDGEMENTS I would like to express my most sincere a nd eternal gratitude to my supervisor Dr. Valerie Harwood for reading my application and seeing someone she wanted to work with. I thank her for believing in me, for her guidance, advice and patience. I would also like to thank the members of my committee, Dr. Angelo DePaola, Dr. James Garey and Dr. Daniel Lim, for all their time, insight and guidance. Special thanks to my parents (George and Scharmaine Gordon) for their support, and the sense of responsibility and discipline they have ins tilled in me, without which I would never have been able to accomplish th is goal. To the members of my immediate and extended family, thank you for keeping tabs on me and making sure I was ok when so far away from home. To the members of my lab go my gratitude for being my sounding board, for all their help with experi ments, their proof reading skills, and for helping keep me focused throughout the years. Jason, thank you for keeping me calm, focused and motivated. Lastly, but by no means least, I would like to express my gratitude to the teachers I have had at Immaculate Conception Prep aratory School, Immaculate Conception High School and the University of the West Indi es, Mona. Even though they may never read this work, they gave me the necessary tools and planted the seeds of love for science that led me to graduate school and ul timately to complete this degree.

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i TABLE OF CONTENTS LIST OF TABLES.............................................................................................................iv LIST OF FIGURES.............................................................................................................v ABSTRACT....................................................................................................................... vi BACKGROUND.................................................................................................................1 Vibrio vulnificus and Disease..................................................................................1 Classification...........................................................................................................2 Occurrence..............................................................................................................3 Virulence Models....................................................................................................4 Animal Models............................................................................................4 Cell Culture Models....................................................................................5 Commonly Studied Viru lence Factors....................................................................7 Capsular Polysaccharide.............................................................................7 Iron Acquisition..........................................................................................8 Flagella......................................................................................................10 Hemolysin/Cytolysin................................................................................11 Virulence Associated Genes.................................................................................13 16S rRNA..................................................................................................13 Virulence Correlated Gene.......................................................................14 Virulence Factors in Other Vibrio spp..................................................................15 Real-Time PCR.....................................................................................................16 Vibrio vulnificus Genomic Fingerprinting............................................................18 References.............................................................................................................22 REAL-TIME PCR ASSAYS FOR QUANTIFICATION AND DIFFERENTIATION OF V. VULNIFICUS STRAINS IN OYSTERS AND WATER.......................................................................................................................32 Abstract.................................................................................................................33 Introduction...........................................................................................................34

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ii Methods.................................................................................................................35 Bacterial Strains........................................................................................35 Preparation of Pure Cultures.....................................................................37 Total Cell Counts......................................................................................37 Primer Development.................................................................................38 Real-Time PCR Protocol..........................................................................38 RFLP Confirmation..................................................................................39 Sensitivity.................................................................................................39 PCR Detection of Native V. vulnificus in Oyster Homogenates...............40 Statistical Analysis....................................................................................41 Results...................................................................................................................41 RFLP Confirmation..................................................................................44 Sensitivity.................................................................................................44 PCR Detection of Native V. vulnificus in Oyster Homogenates...............46 Typing of Clinical and Environmental Isolates........................................46 Discussion.............................................................................................................48 Sensitivity.................................................................................................48 A:B Ratios and Geographic Distribution..................................................49 Acknowledgements...............................................................................................51 References.............................................................................................................51 BOX-PCR GENOTYPING AND MULTILO CUS ANALYSIS OF VIRULENCE ASSOCIATED GENES OF ENVIRONMENTAL V. VULNIFICUS ISOLATES FROM PERMITTED AND PROHIBITED SHELLFISH HARVESTING AREAS..............................................................................................55 Abstract.................................................................................................................56 Introduction...........................................................................................................57 Methods.................................................................................................................61 Bacterial Strains........................................................................................61 Multilocus PCR Analysis..........................................................................63 BOX-PCR Genotyping.............................................................................66 Results and Discussion.........................................................................................67 Multilocus Typing.....................................................................................67 BOX-PCR Genotyping.............................................................................70

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iii References.............................................................................................................82 RESEARCH SIGNIFICANCE..........................................................................................86 Virulence Typing Methods...................................................................................86 Correlations with Virulence and Water Quality...................................................88 References.............................................................................................................90 ABOUT THE AUTHOR.......................................................................................End Page

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iv LIST OF TABLES Table 1. V. vulnificus strains, origin, and type.................................................................43 Table 2. Genotypes of V. vulnificus isolates from oysters, wate r and clinical sources...44 Table 3. Harvest area location and harvest date information along with results of multilocus analysis of isolates from permitted shellfish harvesting areas.........63 Table 4. Harvest area location and harvest date information along with results of multilocus analysis of isolates from pr ohibited shellfish harvesting areas........64

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v LIST OF FIGURES Figure 1. Example of gel showing type A and type B V. vulnificus 16S rDNA PCR products...................................................................................................68 Figure 2. Example of gel showing a) vcgC and b) vcgE gene products..........................68 Figure 3. Example of gel showing viuB gene products...................................................69 Figure 4. Examples of gels showing results for HlyIII PCR attempts.............................69 Figure 5. Agarose gel #1 showing V. vulnificus BOX-PCR patterns..............................71 Figure 6. Agarose gel #2 showing V. vulnificus BOX-PCR patterns..............................71 Figure 7. Agarose gel #3 showing V. vulnificus BOX-PCR patterns..............................72 Figure 8. Agarose gel #4 showing V. vulnificus BOX-PCR patterns..............................72 Figure 9. Agarose gel #5 showing V. vulnificus BOX-PCR patterns..............................73 Figure 10. Dendrogram showing the similarity of type A V. vulnificus from permitted and prohibited she llfish harvesting areas.........................................74 Figure 11. Dendrogram showing the similarity of type A and type AB V. vulnificus from permitted and prohibited shellfish harvesting areas...............76 Figure 12. Dendrogram showing the similarity of type B V. vulnificus from permitted and prohibited she llfish harvesting areas.........................................78 Figure 13. Dendrogram showing the similarity of type B and type AB V. vulnificus from permitted and prohibited shellfish harvesting areas...............79 Figure 14. Dendrogram showing the similarity of all V. vulnificus isolates......................80

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vi MULTILOCUS VIRULENCE TYPING OF CLINICAL AND ENVIRONMENTAL VIBRIO VULNIFICUS ISOLATES KATRINA V. GORDON ABSTRACT The bacterium Vibrio vulnificus is an autochthonous inhabi tant of estuarine waters and also found in shellfish such as oysters. It is a human pathogen of importance in the seafood industry, and can also infect recreational water users. Currently, recognized methods of detection rely upon isolation of pur e cultures which requires at least 24 hours. To reduce the time needed for identification of the pathogen and simultaneously ascertain the virulence potential of the strains present, real-time PCR assays and sample processing procedures were developed (Chapter 1). These assays discriminate between type A (environmental, generally lower virulence) and type B (clinical, higher virulence) isolates. The genetic relationships between environmental V. vulnificus strains isolated from permitted and prohibited shellfish harv esting areas was determined using BOX-PCR genomic fingerprinting coupled with sequen ce analysis of three proposed virulence markers: (1) the virulence correlated gene (vcg ), (2) 16S rRNA type and (3) presence/absence of the vulnibactin gene ( viuB) (Chapter 2). The real-time PCR assays were able to detect the presence of seeded V. vulnificus in environmental water at a concentration of 160 cells 100ml-1. In seeded oyster homogenates, the assays were ab le to detect a minimum of 103 cells and 102 cells per reaction of type A and type B respectively.

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vii The phylogenetic analysis separa ted the majority of type A/ vcgE strains isolated from permitted shellfish harvesting areas from those isolated from prohibited harvesting areas. The genomic (BOX-PCR) fingerprints of type A and type AB isolates were more similar to one another than to t ype B isolates. On ly one type A/ vcgE isolate contained the viuB gene; however, eight type B/ vcgC isolates had that gene. No obvious grouping was discerned between type B/ vcgC isolates from permitted versus prohibited shellfish harvesting areas or between those possessing the viuB gene versus those lacking viuB. These data provide insight into th e ecology and correlation between population biology and general water quality, as gauged by the classification of the shellfish growing area waters. The 16S typing assays can be used for routine rapid t yping to aid in risk assessment and reduce infection frequenc y through consumption of contaminated seafood.

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1 BACKGROUND Vibrio vulnificus and Disease Vibrio vulnificus is a rod shaped, Gram-negativ e opportunistic pathogen which is an autochthonous inhabitant of estuarine and marine waters. Infections can occur due to consumption of raw or undercooked seafood, primarily molluscan shellfish such as oysters (20). Ingestion of the bacterium in healthy individual s is usually harmless but can lead to mild symptoms such as gastroenteri tis. However, life threatening symptoms like septicemia can occur in the chr onically ill, especially those with liver disease. Mortality rates in these cases can be as high as 50% (44, 96).Wound infections are also possible due to trauma associated with handling contaminated seafood or the contact of open wounds with water containing V. vulnificus Wound infections can become so severe that amputation is necessary to stop the spread of infection (68). Millions of individuals throughout the world eat raw oys ters and come into contact with waters containing V. vulnificus, yet the rate of infection is very low [less than 1 person in a million without predisposing illnesses (12)]. Predisposing illnesses include liver cirrhosis, hemochromatosis and other immune-suppressing illnesses such as AIDS (26, 96). A number of antibiotics have been reported as effective for infection treatment based on in vitro susceptibility tests. These include te tracycline (9), ampicillin and broad spectrum cephalosporins (18) and a combination of cefotaxime and minocyc line. Most recently, treatment with doxycycline has been advocated (39).

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2 One study investigating V. vulnificus infections between 1988 and 1996 found that 422 persons contracted infections in the USA (87). In Florida, V. vulnificus infections have been noted as the leading cause of food-related illnesses (47). Between 2000-2007, an average of 28 cases of V. vulnificus infections were confirmed in Florida with the highest number of confirmed infections, 45 cases, reported in 2003 (MERLIN disease reporting systemhttp://www.floridacharts.com/ merlin/freqrpt.asp). In 2007 there were 21 confirmed cases of V. vulnificus infections in Florida. V. vulnificus is also becoming an important pathogen throughout the USA as noted in a recent Morbidity and Mortality Weekly Report (MMWR) surveillance summery of recreational water associated diseases and outbreaks. Among the reported V. vulnificus infections, 82.7% required hospitalization and 12.8% were fatal, whic h were the highest rates among the Vibrio spp. monitored (24). Classification Generally, the biochemical characteristics of this organism include its ability to produce gelatinase, lysine decarboxylase and ox idase enzymes but not urease; ability to ferment D-cellobiose, D-mannose, lactose and ONPG (o-nitrophenyl-beta-Dgalactopyranoside) and inability to ferment sucrose, arabinose and butanediol (negative Voges-Proskauer reaction) (25, 99). Variations in strain biochemical profiles and host specificity have lead to the separation of this bacterium into three biotypes. Biotype 1 is indole and ornithine decarboxylase positive. Biotype 1 strains exhibit a variety of lipopolysaccharide (LPS) types, are pathogenic to humans and are commonly found in shellfish. V. vulnificus biotype 2 is indole and ornithine decarboxylase negative. All

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3 Biotype 2 isolates exhibit a similar LPS t ype (serogroup E) and are normally responsible for infections of eels and marine invertebrates (4, 95). Biotype 2 V. vulnificus have however, been isolated from infected humans (1 ). A third, more recently classified type is biotype 3. Biotype 3 strains are citrate, D-cellobiose, lactose and ONPG negative (13) and are mainly associated with fish, although se vere human infections have been reported in Israel (5). Biotype 1 V. vulnificus will be the focus of this research. Occurrence There is a well known folk saying that warns against consuming raw oysters during months without the lett er r (May, June, July, Augus t). Research has shown that this long held belief that consumption of oysters harvested during these months is bad for you holds some scientific merit. Temperat ure and salinity have been shown to play a great part in the surviv al (or culturability) of V. vulnificus and therefore may influence the incidence of infections caused by this pathogen. Moreover, in a study by Motes et al (72), it was found that V. vulnificus concentrations in oysters were influenced most greatly by water temperature. Temperatures below 26C caused a rapid decline in V. vulnificus concentrations (from 104 MPN/ g to <10 MPN/ g in Gulf Coast oysters) while above this temperature, concentr ations remained constant (104 MPN/ g). Tamplin and Capers (97) actually detailed growth of V. vulnificus in and on oysters and release of the bacterium into the surrounding water. Us ing MPN enumeration techniques, they calculated that between 105 to 106 V. vulnificus were released each hour when the water temperature was kept at 21C. Conversely, salinities of 5 to 25 ppt led to increased

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4 densities (>103 organisms/g) while above 25 ppt densities decreased to <102 /gram (72). Other studies have routinely found concentrations of 103 organisms/g (47, 107). Virulence Models Animal models. In light of the relatively low numbers of individuals becoming ill after eating oysters containing V. vulnificus [0.2 per million persons in Florida, with 3 million Floridians consuming raw oysters (12)], it seems that there is some variation in strain virulence and or host suscepti bility. The high level of mortality of V. vulnificus infections in humans makes dose-response studies unethical Therefore, in order to determine any strain-dependent variation in virulence of V. vulnificus isolates, an animal model of infection is the most useful strategy (44, 96). Du e to the observation that the majority of infections occur in persons who are immunocompromised or have diseases causing elevated iron concentrations (11, 12), and because iron had previously been shown to increase the 50% lethal dose (LD50) of other pathogens ( 45), an iron-overloaded model of infectivity has been developed. This work began with a study conducted by Wright et. al in 1981 (111), where intraperit oneal (ip) inoculation of mice with ferric ammonium citrate was used to mimic the high iron content found in humans that are susceptible to infection. Using one strain (CDC C7184) to challenge mice orally, they observed a 6-log reduction in the LD50 (from 106 to 1.1 CFU). This work was continued by Stelma et al. (94) in a study comparing the LD50 reduction for environmental and clinical V. vulnificus isolates, in which iron-overload was simulated by treating mice with ferric ammonium citrate and iron dextran. They concluded that ir on overload using irondextran was a more reliable model compared to ferric ammonium citrate and was capable

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5 of differentiating even weakly-virulent is olates from avirulent ones. The authors theorized that this was due to the ability of the iron-dextran to persist longer in the bloodstream than ferric ammonium citrate. This study also classified is olates into virulent and avirulent categories based on the extent of reduction in LD50 in untreated vs. ironoverloaded mice. Virulent strains were t hose which experienced a 3.5 log-reduction. The iron-dextran induced, iron-overl oaded mouse model is now wide ly used to investigate the virulence of V. vulnificus isolates and the effects of the many virulence factors being studied (47, 53, 60, 83, 92, 93) even though nearly all V. vulnificus strains regardless of source appear to virulent in this model. Cell culture models. Cell culture has been used fo r virulence testing in a limited capacity, mostly to observe the effect of toxins (55, 61) and to monitor host cell interactions (83). One early virulence study of the hemolysin/cytolysin toxin by Kreger and Lockwood (61) used mouse, guinea pig, ra bbit and human erythrocytes to determine the toxin concentration necessary to lyse 50% of the erythroc yte preparation. They also used the Chinese hamster ovary cell line to determine the minimum cytotoxic dose for different preparations of the toxin (61). Chin ese hamster ovary cells have been used in several subsequent studies (36, 110) as we ll as erythrocytes fro m several different animals (36). Several studies have suggest ed that erythrocyte lysis occurs through cholesterol receptor-mediated pore formation in the cell membrane (36, 55, 89). In vivo-induced antigen technology (IVIAT) was used to create a clone library containing genes expressed by V. vulnificus in septicemia patients (57). To isolate the genes expressed during infection, the IVIA T method used cultures of a clinical V. vulnificus strain grown in vitro to absorb and remove antibodies that were specific for

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6 proteins expressed in free-living V. vulnificus from the pooled serum of three infected individuals. The remaining antibodies, wh ich were presumably unique to proteins expressed by V. vulnificus during infection, were hybridized to the proteins generated from an expression library of the clinical strain. The twelve clones with which antibodies hybridized were studied further, and repres ented products from genes including a putative serine/threonine kinase, transcriptional ac tivator genes (IlvY and HlyU), a putative lipoprotein (YaeC) as well as biosynthesis/me tabolism genes (PyrH, PurH, and IlvC) and secretion genes (TatB and plasmid Achromobacter secretion (PAS) fact or). These genes were then mutated in clinical V. vulnificus strains. The HeLa cell line (cancer cells) was then used to study the change in cytotoxicity due to the lack of that gene product in each mutant. The LD50 of each strain was also determin ed using ip inoculation of the V. vulnificus mutants in mice. The mutants for the hlyU pyrH and purH genes all showed decreased cytotoxicity and increased LD50, indicating that these ge nes may be involved in virulence (57). Studies have also been conducted in wh ich animal or human cell lines were exposed to intact, viable V. vulnificus cells. Paranjpye et al used the human epidermoid carcinoma cell line (HEp-2) to study host cell adhe rence (83). Kashimoto et al (52) used both mice and the macrophage-like cell line (J774) to investigate the ability of clinical and environmental isolates to cause apoptosis. Both models gave the same result; the clinical isolate was seen to cause apoptosis in host cells while the environmental strain used in both models failed to cause apoptos is (52). These culture d cell models can be used to observe more directly how different toxins and mutations in various genes affect

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7 target cells, however, to observe the eff ect on the organism, and in a more dynamic setting, animal models ar e more appropriate (38) Commonly Studied Virulence Factors Capsular polysaccharide. The capsular polysaccharide (CPS) is among the most important virulence factors of V. vulnificus (38) as it gives prot ection from the hosts immune response (68, 95, 114). The important association between the presence of a capsule and the virulence of V. vulnificus was first observed by Yo shida et al (114). Electron microscopy on cells stained with ruthenium red was used to observe the presence/absence of the CPS layer for opaque and translucent types respectively, of four strains. Opaque, capsulated strains had an obvious CPS layer by electron microscopy and generally lower LD50 compared to translucent, unencapsulated strains (assessed by injection into mice). This study also noted th e ability of strains to transition between opaque and translucent type colony mor phology, and were able to determine the frequency of the phase shifts for two stra ins where the opaque to translucent shift occurred more frequently. Simpson et al (91) obtained similar results with electron microscopy and LD50 measurements in mice; however they did not observe reversion of translucent types back to opaque. Work on the CPS continued with the production of the first translucent transposon mutants by Wright et al (112). These mutants were unable to revert to the opaque type after passage th rough iron-overloaded mice, while the naturally translucent strain could revert to being opaque. Perhap s most importantly, Zuppardo and Siebeling were able to apply the molecular version of Kochs postulates (37) when they created a

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8 mutant incapable of producing a CPS layer. This mutant was nonlethal in the ironoverload mouse model, and lethality was re stored after complementation (115). This experiment confirmed the CPS layer as a virulence factor of V. vulnificus Iron acquisition. As previously noted, V. vulnificus causes serious infections in hosts with increased serum iron concentrations. In fact it is among only a small number of opportunistic pathogens that have limite d iron acquisition capability (106). While experiments with iron-overloaded mice readily confirmed that increased iron availability leads to decreased LD50 (94, 111, 112), pinpointing the proteins involved in iron acquisition would prove more challenging. Simpson and Oliver were among the early investigators into the proteins involved in iron scavenging in V. vulnificus (90). By growing V. vulnificus in iron-limited medium they were able to induce the production of phenolate (or catechol-like) and hydroxamate cl asses of siderophore in virulent strains while the avirulent st rain they studied did not produ ce the phenolate siderophore. These molecules are used to chelate iron from the organisms surroundings and transport it across the cell membrane, where the iron is then stripped from the prot ein and used in the cell (105). Helms et al. found that the intraper itoneal (ip) injection of hemoglobin, methemoglobin and hematin would increas e the lethality of ip-administered V. vulnificus in mice (41); concluding that these molecu les are sources of Fe for growth during infection. In 1994, Okujo et al we re able to characterize the structure of the catechol-like siderophore, vulnibactin (79). Later, they would continue their investigations into this siderophore to find that iron ut ilization probably occurs in two steps. The iron-carrying host proteins, transferrin and lactoferrin are first cleaved by an extracellular protease into

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9 smaller peptide fragments from which th e vulnibactin sidero phore can more easily acquire bound iron (78). Using a mutant incap able of utilizing transferrin-bound iron, Litwin et al. (69) were able to elucidate the sequence of the genes venB and viuB (vulnibactin). These genes are involved in the production of the catechol siderophore implicated in the utilization of iron from transferrin. Although other factors are undoubtedly involved in the iron acquisition mechanisms of V. vulnificus the statistically significant increase in the LD50 of the mutant in that study again showed the involvement if the catechol sider ophore and these two genes in virulence. More recently, in an attempt to develop sc reening tests for highly virulent strains of V. vulnificus Panicker et al developed primers to amplify the viuB gene and incorporated them into a multiplex PCR assay (81, 82). The assay was able to specifically detect the presence of V. vulnificus as well as determine their putative virule nce potential (based on the presence or absence of the vulniba ctin gene). Detection levels as low as 1 CFU V. vulnificus g-1 oyster homogenate were achieve d when DNA was extracted from seeded oyster homogenates after overnight enri chment. These primers were also used in this dissertation work to help determine the relationship among virulence factors, genomic type and environment among V. vulnificus strains. Several subsequent studies have delved deeper into the presence, absence and expression of the viuB gene in clinical and enviro nmental strains and under varying environmental stresses (8, 50, 80). Bogard and Ol iver (8) showed that both clinical and environmental strains that possessed the viuB gene were able to survive for significantly longer periods of time in human serum than those strains without the gene. When they increased the iron concentration in the serum by adding ferric ammonium citrate, strains

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10 with and without the gene were able to survive for essentially the same period of time. In this study they also investig ated the effect of osmotic/ nutrient downshifts by incubating strains in artificial seawater to simulate environmental c onditions for various time periods before inoculating them into human serum. By imitating the changes experienced by the strains during infection, they were able to show that the vcgC type strains (all containing the viuB gene) showed a statistically signifi cant increase in survivability over the environmental strains (lacking the viuB gene). These two findings suggest that the increased survivability of vcgC viuB + strains in human serum, and by extension in the human host, account for these strains being more virulent and most frequently isolated from infected individuals (8). Jones et al (50) st udied the expression of the viuB gene under high (31 ppt) and moderate (21 ppt) salin ity. They noted that expression ceased in as little as 30 min in the moderate salinity microcosm and theorized that this might be due to the role of viuB in pathogenesis, while other gene s are involved in scavenging iron when the organism is free-living in the aquatic environment. Flagella. Vibrio vulnificus are motile via a polar flagellum (3). Using marinerbased transposon mutagenesis (Tn Himar1 ) in an effort to identify additional virulence factors of V. vulnificus, Kim and Rhee 2003 (58) stumbled upon a portion of the flg operon, which encodes the flagella r basal body rod proteins. The Tn Himar1 had inserted into the flgC gene of the flgBCDE operon, interrupting the comp letion of biosynthesis of the flagellar basal body. This mutation resulted in a significant decrease in motility, cytotoxicity and adhesion to HeLa cells as well as an increased LD50 for the mutant V. vulnificus strain in suckling mice when the bacteria were inoculated directly into the stomach (58).

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11 This study served as a starting point for i nvestigation of the role of flagella in virulence (58). Building on this resear ch, Lee et al. (63) constructed a flgE knockout V. vulnificus mutant which was not able to produce flagella and showed increased LD50. Complementation was able to restore the ab ility to produce flagella in pure culture. However no observable decrease in LD50 by the motile, complemented V. vulnificus compared to the knockout mutant was seen in challenged mice. Th e authors propose that this conflicting result is due to loss of the complementation plasmid after inoculation because of the lack of antibiotic selective pressure in mice (63). Due to the number of genes involved in the production of flagella, no molecular assays targeting these genes have been de veloped to date to identify virulent vs. attenuated strains of V. vulnificus. However, one group has gone as far as to investigate the usefulness of flagellin genes ( flaA flaB flaF flaC flaD and flaE ) in the production of a mucosal vaccine (64). They were able to show that intranasal administration of FlaB in mice was able to effectively produce a mu cosal and systematic immune response and therefore has the potential to induce protective immunity (64). Hemolysin/Cytolysin. The hemolysin/cytolysin toxin is among the most thoroughly investigated extracel lular protein products of V. vulnificus. Research dates back to 1981 when both Kreger and Johnson re ported detection of hemolytic activity in V. vulnificus cultures (48, 61). Extracts of V. vulnificus cultures were used to demonstrate cytolytic activity against red blood cells as well as Chinese hamster ovary (CHO) cells (61). They also found that more virulent strains (LD50 of 56 versus 29 CFU for their weakly virulent strain) produced a higher tite r of toxin, leading to hypotheses about the role of this toxin in virule nce potential and pathogenesis.

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12 Continued research on the cytolysin w ould see several papers published in 1985 detailing the elucidation of the amino aci d composition and sequence of the first 10 amino terminal residues (36), analysis of th e mode of action (89), and cloning of the cytolysin gene (vvhA ) (110). The first rapid assays for V. vulnificus identification, involving indirect immunofluor escence (IIF) (34) or a DNA-probe based on the sequence of the cytolysin gene (110) were also deve loped. The ability to use the DNA probe to detect all V. vulnificus strains regardless of source (clini cal or environmental) led to the question of whether the toxi n was actually expressed during infection. Although Wright et al (110) noted the expression of the toxin in vitro for a number of clinical and environmental isolates, research continued on determining whether the toxin is produced, conditions under which it is produced as we ll as the actual role of the toxin during infection (32, 33, 35, 55, 62, 66, 109, 113). Notable discoveries included that of Wright and Morris (109), that cytolysin-deficient mutants had the same LD50 as their cytolysinproducing parent strains in mice. These experi ments led to the conclu sion that this toxin cannot be used as an absolute judge of vi rulence. The nucleic acid sequence of the vvhA gene was also elucidated (113). Today the Food and Drug Administration (FDA) advocates the use of a hemolysin/cytolysi n oligonucleotide probe (108) for use in identification and enumeration of the total V. vulnificus in seafood (54). Although many years of research effort have focused on the vvhA -encoded hemolysin, two other less studied hemolysins, vllY (14) and hlyIII (17) have been cloned and sequenced. Chen et al discovered th e latter hemolysin most recently, in 2004. hlyIIIdeficient mutants showed increased LD50 compared to their parent strains, making this

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13 gene another promising piece of the virulence puzzle of V. vulnificus (although no additional research has been published to date. Virulence Associated Genes In addition to the above mentioned viru lence factors, the variation in the sequences of at least two other genes (the 16S rRNA and vcg ) have also been correlated with, though not directly related to, virulence. 16S rRNA. Aznar et al (2) first noted the 17 nucleotide variation in the sequence of the 16S rRNA gene of V. vulnificus isolates which separates them into two groups, denoted type A and type B. The authors we re able to construct oligonucleotide probes targeting the 16S rDNA, and a hybridization strategy that was able to differentiate between the two types (2). In 2001, Kim and Jeong developed a tri-primer PCR assay to differentiate between type A and type B V. vulnificus 16S rDNA sequences. Their analysis of 40 environmental V. vulnificus isolates (from water, sediment and oysters) showed 35% of the isolates to be type A a nd the majority to be type B (56). However, later research by Nilsson et al (73) employi ng a terminal restriction fragment length polymorphism (T-RFLP) assay util izing the presence/absence of Alu I and Hae III restriction sites in a 492 bp re gion of the 16S rDNA to differentiate the two types would show conflicting result s. They tested 33 V. vulnificus isolates of environmental origin (isolated from retail oysters) and 34 V. vulnificus isolates from clinical origin (isolated from patients with primary septicemia infect ed through ingestion of raw oysters). Their results showed that the type A V. vulnificus were in the majority in environmental isolates (94%) and that type B V. vulnificus were in the majority in clinical isolates (76%)

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14 especially fatal clinical infections (94%). This implied that type B V. vulnificus were most often responsible for human infections, perhaps due to a highe r virulence potential than type A. A TaqMan real-time PCR assa y was then developed to allow more rapid V. vulnificus strain typing (102). Using essentially the same set of V. vulnificus isolates, the same proportion of type A in environmental isolates and type B in clinical isolates was observed as in the Nilsson study. In an e ffort to improve on this real-time PCR assay, our lab designed two V. vulnificus specific, SYBR Green mediat ed real-time assays for type A and type B (31). These assays bypass V. vulnificus culturing and DNA extraction steps, enabling more rapi d analysis of oyster homogenates and water samples. Virulence correlated gene. The virulence correlated gene ( vcg ) was first discovered in 1999 as a ~200 bp band in ra ndomly amplified polymorphic DNA (RAPD) PCR fingerprints that was presen t in most clinical and few e nvironmental isolates (104). This lead to the theory that the presence if this band in an isolate may give some information about its virulence. Building on th is finding, Rosche et al. (85) sequenced a 1700bp amplicon containing this 200 bp target fr om several clinical and environmental V. vulnificus isolates. They determined that clinical and environmental is olates had differing sequences (and were termed the vcgC and vcgE genes) in this ~200 bp region and were able to create PCR primers to diffe rentiate the two. PCR analysis of 55 V. vulnificus isolates using these new primers found 93% of environmental isolates had the vcgE sequence while 72% of c linical isolates were vcgC This typing strategy has been used in several studies as a means of indi cating the level of virulence of V. vulnificus isolates (8, 15) as well as in this work.

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15 Warner and Oliver also used this ty ping strategy in 2008 to study the proportions of vcgC and vcgE V. vulnificus strains present in oysters and their overlying waters (103). They found that 84.4% of th e oyster isolates had the vcgE (environmental) gene sequence while in the water column there was an almo st even distribution of the two types. As water temperatures increased, the relative proportion of the vcgC type increased in both the oysters and water column. These data point to a preferential con centration of one type of V. vulnificus in oysters and help explain why so few people become infected. These studies also highlight the usef ulness of this typing method to investigate the virulence potential of V. vulnificus strains. Virulence Factors in Other Vibrio spp. Vibrio cholerae and Vibrio parahaemolyticus are closely related to V. vulnificus (16) and are also important hum an pathogens. The virulence of V. cholerae is primarily dictated by its ability to produ ce a heat labile enterotoxin, called the cholera toxin (CT). The toxin is produced after colonization of th e epithelial cells of the small intestine. Secretion of the CT results in profuse watery diarrhea that is characteristic of cholera (84). Like V. vulnificus V. parahaemolyticus produces several hemolysins (five have been identified so far (67)). Studies have show n that two of these pl ay an important role in virulence (i) a thermally st able direct acting hemolysin ( tdh ) and (ii) a thermally stable direct actingrelated hemolysin (trh ) (74-76). These genes can be targeted to determine the presence of pathogenic V. parahaemolyticus strains (7, 21, 77). Another hemolysin, the thermolabile hemolysin ( tlh), is present in all V. parahaemolyticus strains and is used

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16 in assays for detection of the total V. parahaemolyticus population (77) in much the same way the cytolysin gene is used to detect V. vulnificus (65). V. cholerae also produces a hemolysin-cytolysin protein; however its ro le in virulence is still uncertain (51). Real-Time PCR It could be said that modern molecular biology began in 1983 with the creation of polymerase chain reaction (PCR) by Kary B Mu llis. In 1985, Saiki et al (86) made the first published mention of the procedure. Over the next two decades, it has been modified and improved upon to become a mainstay in any serious molecular laboratory. One of these improvements added the ability to mon itor the amplification of DNA as the reaction is progressing. This new form of PCR has b een termed kinetic PCR, quantitative PCR or real-time PCR. One of the first published accounts of this new type of PCR is from Higuchi et al (43) using ethidium bromide (EtBr) to bind dsDNA, UV lamps to excite the bound EtBr and a camera to capture fluorescence after each annealing/extension step. Today, other dsDNA binding dyes such as SYBR Green are us ed (31) and the equipment necessary has morphed into single units capable of both amp lification and detection. Perhaps the single greatest advantage of this method is its cost effectiveness compared to newer methods utilizing fluorogenic probes. Molecular beacons were developed in 1996 (100) giving birth to a second method of real-time PCR. These beacons are essent ially oligonucleotides having a middle portion complementary in sequence to the target to be amplified, and ends which are complementary to each other and not the target so the beacon forms a stem-and-loop

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17 structure. At each end of the oligonucleot ide is a fluorescent and a non-fluorescent, quenching moiety. When not hybridized to target DNA, the stem-loop shape keeps the two ends so close that the quencher prohibits the fluoresce nt moiety from fluorescing when excited during annealing/extension. Howe ver, when hybridized to the target, the increased distance between the two allo ws the fluorophore to fluoresce (100). Yet another assay involves the use of probes (termed TaqMan probes) which are similar to molecular beacons but are cleaved during elongation by the 5'-3' nucleolytic activity of the DNA polymerase. The release of the reporter allows it to be sufficiently far from the quencher to allow detection of its excitation emissions (27, 40). With all these methods, a sigmoid curve is ideally pr oduced during amplification protocols. The cycle at which logarithmic amplification occurs can then be used to determine the amount of target added to the initial reaction by comparing with known standards (40). Today these real-time PCR reaction me thods are routinely used to detect presence/absence as well as quantify various targets in well funde d laboratories around the world. There are obvious advantages of real-time PCR over conventional PCR, including the ability to follow reactions as they are happening instead of having to wait until reaction completion and further time consuming manipulations. Additionally, molecular beacon and TaqMan probe mediat ed reactions have essentially three oligonucleotides binding to the target (primers and probe), which adds another level of stringency over dsDNA-binding mediated r eactions using only tw o oligonucleotides (primers). Use of an internal control wh ich allows the determination of any PCR inhibition in each sample is also gaining momentum (77).

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18 Vibrio vulnificus Genomic Fingerprinting Various methods have been employed in an attempt to differentiate V. vulnificus strains and group them on the basis of char acteristics such as geographic origin or virulence potential, which some would argue, both go hand in hand for a majority of isolates. One group (10) used clamped homogenous electric field (CHEF) gel electrophoresis on genomic DNA digested with rare cutting restriction enzymes to type 95 V. vulnificus isolates from three oysters. They found 60 different patterns with Stf I and 53 patterns with Srf I, showing a high degree of gene tic diversity among the isolates. Tamplin followed this work with an analysis of clinical and environmental isolates by restriction fragment length polymorphisms (RFLP) separated by pulse field gel electrophoresis (PFGE) as well as ribotyping (98). Again, a hi gh degree of diversity was observed among all the isolates tested, and the le vel of similarity between isolates varied based on the typing method used. Ribotyping show ed the highest degree of similarity (90%) between isolates from similar isolati on environments, for 25 clinical isolates and 28 environmental isolates while the highest si milarity for PFGE was between two clinical isolates (92%) (98). PFGE was later used to type V. vulnificus isolated from the blood of infected patients in addition to oysters implicated in causing the infections when possible (47). This group was able to show that, alt hough there were several st rains present in the infecting oysters, all V. vulnificus isolated from the patient were of the same strain (47). This was a very significant finding alludi ng to differential viru lence potential among strains and supporting the need to determine a method for detec tion of strains with greater virulence potential.

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19 Other methods recently employed for V. vulnificus typing include multilocus sequence typing (MLST) (6, 70) and direct genome restriction enzyme analysis (DGREA) (30). MLST involves sequence analysis of internal portions of housekeeping genes. There are some advantages of a sequence based method such as this over electrophoresis based methods such as PFGE. Pe rhaps the most significant of these is the fact that the sequences obtained with MLST can be easily shared and compared between laboratories (70). MLST as outlined for V. vulnificus (6) involves comparison of the sequence variation in a 324-480 bp portion of 10 housekeeping genes, five from chromosome 1 ( glp encoding glucose-6-phosphate isomerase; gyrB encoding DNA gyrase-subunit B; mdh encoding malate-lactate dehydrogenase; metG encoding methionyl-tRNA synthase and purM encoding phosphoribosylaminoimidazole synthetase) and five from chromosome 2 ( dtdS encoding threonine dehydrogenase; lysA encoding diaminopimelate decarboxylase; pntA encoding transhydrogenase alpha sununit; pyrC encoding dihydroorotase and tnaA encoding tryptophanase). Using MLST, Bisharat et al. was able to determine that bi otype 3 isolates from an outbreak in Israel are a hybrid genotype of biotype 1 and 2 V. vulnificus Later analysis of these MLST profiles by the same group showed that recombin ation, rather than mutation, was most responsible for the strain diversity se en between isolates (6). The online V. vulnificus MLST database (http://pubmlst.org/vvulnificu s/) was used to access MLST profiles for a comparison between PFGE and direct genome restriction enzyme analysis (DGREA) (30). DGREA uses restriction en zymes such as the six cutter NaeI to digest genomic DNA, which is then electrophoresed on a non-denaturing polyacrylamide gel. The researchers found that DGREA was able to type a larger number of the strains selected

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20 for the study, the clusters formed agreed mo re frequently with the MLST and other typing data available, and would be a fast er more cost effective typing method than PFGE (30). Several short repetitive DNA sequences have been identified in many prokaryotic species. These include palindromic units (PU) (29), repetitive extragenic palindromes (REP) (22, 28, 42) and enterobacterial repetitiv e intergenic consensus (ERIC) sequences (46, 88). Soon after their identif ication, these repeat sequences found use as targets for fingerprinting techniques a nd bacterial classification (19, 101). BOX elements are another class of repe titive sequences which were discovered in Streptococcus pneumoniae These elements were initially found to be composed of three types, boxA, boxB and boxC (71). In 1995, after having deve loped several primers for the box A, B and C subunits, Koeuth et al found the box A el ement to be the most conserved between bacterial species and that it was possible to have only one type of box subunit (59). These primers have since been used in several studies to classify bacteria in microbial source tracking (23, 49). Other fingerprinting techni ques have been used to investigate the diversity of V. vulnificus genotypes in oysters (47) and to investigate the relationships between genotypic patterns and other virulence associated genes (15). Chatzidaki-Livanis et al combined rep PCR typing with multilocus PCR analysis of 33 clinical V. vulnificus isolates and 35 environmental isolates (mos t of them from retail oysters). The loci targeted included the 16S rRNA, vcg and CPS operon. The rep-PCR patterns separated the isolates into 7 clusters. The majority of cluster I and 100% of cluster VII were composed of clinical isolates containing the mo re virulent version of all the loci analyzed (16S rRNA type B, vcgC and CPS type 1) (15). These findings underline the ability of

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21 these fingerprinting methods to differentiate is olates based on virulence potential, and the usefulness of multilocus typing to identify relationships among these strains. In this dissertation work, real-time PCR assays were developed to differentiate type A and type B V. vulnificus Sample processing protocol s were then developed to allow testing of water samples without culturing, and oyster homogenates without necessitating the isolation of V. vulnificus colonies. These methodologies were extensively tested for their sensitivity a nd specificity. Both these procedures are improvements on current water and seafood testing procedures for this pathogen (54) which will allow determination of the virulence potential of the V. vulnificus present in a much shorter time frame. BOX A2R primers (59) have also been used to determine the genotypic fingerprints of several V. vulnificus isolates from shellf ish harvesting areas of varying water quality. These fingerprints were combined with multilocus typing using the 16S rRNA, vcg and viuB genes to determine the relationship between virulence potential and shellfish harvesting area water quality. The end result of this research is a new testing option for regulatory agencies that could be implemented to afford more rapid assessment of the risk associated with recreational water and seafood. In add ition, the genetic relationships and sequence/presence of virulence associated genes in environmental V. vulnificus isolates has given new insight into the population biology of V. vulnificus strains isolated from various environments.

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22 References 1. Amaro, C., and E. G. Biosca. 1996. Vibrio vulnificus biotype 2, pathogenic for eels, is also an opportunistic pathoge n for humans. Appl Environ Microbiol 62: 1454-7. 2. Aznar, R., W. Ludwig, R. I. Amann, and K. H. Schleifer. 1994. Sequence determination of rRNA genes of pathogenic Vibrio species and whole-cell identification of Vibrio vulnificus with rRNA-targeted oligonucleotide probes. Int J Syst Bacteriol 44: 330-337. 3. Bergey, D. H., and J. G. Holt. 1993. Bergey's manual of determinative bacteriology, 9th ed. Williams & Wilkins, Baltimore. 4. Biosca, E. G., J. D. Oliver, and C. Amaro. 1996. Phenotypic characterization of Vibrio vulnificus biotype 2, a lipopolysaccharide-based homogeneous O serogroup within Vibrio vulnificus Appl Environ Microbiol 62: 918-27. 5. Bisharat, N., V. Agmon, R. Finkel stein, R. Raz, and G. Ben-Dror. 1999. Clinical, epidemiological, and microbiological features of Vibrio vulnificus biogroup 3 causing outbreaks of wound infection and bacteraemia in Israel. Israel Vibrio Study Group. Lancet 354: 1421-1424. 6. Bisharat, N., D. I. Cohen, M. C. Maiden, D. W. Crook, T. Peto, and R. M. Harding. 2007. The evolution of genetic st ructure in the marine pathogen, Vibrio vulnificus Infect Genet Evol 7: 685-93. 7. Blackstone, G. M., J. L. Nordstrom, M. C. Vickery, M. D. Bowen, R. F. Meyer, and A. DePaola. 2003. Detection of pathogenic Vibrio parahaemolyticus in oyster enrichments by real time PCR. J Microbiol Methods 53: 149-55. 8. Bogard, R. W., and J. D. Oliver. 2007. Role of iron in human serum resistance of the clinical and environmental Vibrio vulnificus genotypes. Appl Environ Microbiol 73: 7501-5. 9. Bowdre, J. H., J. H. Hull, and D. M. Cocchetto. 1983. Antibiotic efficacy against Vibrio vulnificus in the mouse: superiority of tetracycline. J Pharmacol Exp Ther 225: 595-8. 10. Buchrieser, C., V. V. Gangar, R. L. Murphree, M. L. Tamplin, and C. W. Kaspar. 1995. Multiple Vibrio vulnificus strains in oysters as demonstrated by clamped homogeneous electric field gel el ectrophoresis. Appl Environ Microbiol 61: 1163-1168. 11. CDC. 1996. Vibrio vulnificus infections associated w ith eating raw oysters--Los Angeles, 1996. Morb Mortal Wkly Rep 45: 621-4.

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23 12. CDC. 1993. Vibrio vulnificus infections associated with raw oyster consumption-Florida, 1981-1992. Morb Mortal Wkly Rep 42: 405-407. 13. Cerda-Cuellar, M., L. Permin, J. L. Larsen, and A. R. Blanch. 2001. Comparison of selective me dia for the detection of Vibrio vulnificus in environmental samples. J Appl Microbiol 91: 322-7. 14. Chang, T. M., Y. C. Chuang, J. H. Su, and M. C. Chang. 1997. Cloning and sequence analysis of a novel hemolysin gene (vllY) from Vibrio vulnificus Appl Environ Microbiol 63: 3851-7. 15. Chatzidaki-Livanis, M., M. A. Hubbard, K. V. Gordon, V. J. Harwood, and A. C. Wright. 2006. Genetic distinctions among clinical and environmental strains of Vibrio vulnificus Appl Environ Microbiol. 72: 6136-6141. 16. Chen, C. Y., K. M. Wu, Y. C. Chang, C. H. Chang, H. C. Tsai, T. L. Liao, Y. M. Liu, H. J. Chen, A. B. Shen, J. C. Li, T. L. Su, C. P. Shao, C. T. Lee, L. I. Hor, and S. F. Tsai. 2003. Comparative genome analysis of Vibrio vulnificus a marine pathogen. Genome Res 13: 2577-2587. 17. Chen, Y. C., M. C. Chang, Y. C. Chuang, and C. L. Jeang. 2004. Characterization and virulence of hemolysin III from Vibrio vulnificus. Curr Microbiol 49: 175-9. 18. Chuang, Y. C., C. Y. Yuan, C. Y. Liu, C. K. Lan, and A. H. Huang. 1992. Vibrio vulnificus infection in Taiwan: report of 28 cases and review of clinical manifestations and treatment. Clin Infect Dis 15: 271-6. 19. de Bruijn, F. J. 1992. Use of repetitive (repetitive extragenic palindromic and enterobacterial repetitive intergeneric consensus) sequences and the polymerase chain reaction to fingerp rint the genomes of Rhizobium meliloti isolates and other soil bacteria. Appl Environ Microbiol 58: 2180-7. 20. DePaola, A., G. M. Capers, and D. Alexander. 1994. Densities of Vibrio vulnificus in the intestines of fish from the U.S. Gulf Coast. Appl Environ Microbiol 60: 984-8. 21. DePaola, A., J. L. Nordstrom, J. C. Bo wers, J. G. Wells, and D. W. Cook. 2003. Seasonal abundance of total and pathogenic Vibrio parahaemolyticus in Alabama oysters. Appl Environ Microbiol 69: 1521-6. 22. Dimri, G. P., K. E. Rudd, M. K. Morgan, H. Bayat, and G. F. Ames. 1992. Physical mapping of repetitive extr agenic palindromic sequences in Escherichia coli and phylogenetic distribution among Escherichia coli strains and other enteric bacteria. J Bacteriol 174: 4583-93.

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24 23. Dombek, P. E., L. K. Johnson, S. T. Zimmerley, and M. J. Sadowsky. 2000. Use of repetitive DNA sequences and the PCR to differentiate Escherichia coli isolates from human and animal sources. Appl Environ Microbiol 66: 2572-7. 24. Dziuban, E. J., J. L. Liang, G. F. Craun, V. Hill, P. A. Yu, J. Painter, M. R. Moore, R. L. Calderon, S. L. Roy, and M. J. Beach. 2006. Surveillance for waterborne disease and outbreaks associ ated with recreational water--United States, 2003-2004. MMWR Surveill Summ 55: 1-30. 25. Elliot, E. L., C. A. Kaysner, L. Jackson, and M. L. Tamplin. 1995. Vibrio cholerae, V. parahaemolyticus V. vulnificus and other Vibrio spp., p. 9.01-9.27, In Food and Drug Administration Bacteriol ogical Analytical Manual, vol. 8ed. Association of Official Anal ytical Chemists, Arlington, VA. 26. Gholami, P., S. Q. Lew, and K. C. Klontz. 1998. Raw shellfish consumption among renal disease patients. A risk factor for severe Vibrio vulnificus infection. Am J Prev Med 15: 243-5. 27. Gibson, U. E., C. A. Heid, and P. M. Williams. 1996. A novel method for real time quantitative RT-PCR. Genome Res 6: 995-1001. 28. Gilson, E., J. M. Clement, D. Brutlag, and M. Hofnung. 1984. A family of dispersed repetitive extragenic palindromic DNA sequences in E. coli. Embo J 3: 1417-21. 29. Gilson, E., D. Perrin, W. Saurin, and M. Hofnung. 1987. Species specificity of bacterial palindromic units. J Mol Evol 25: 371-3. 30. Gonzalez-Escalona, N., B. Whitne y, L. A. Jaykus, and A. DePaola. 2007. Comparison of direct genome restricti on enzyme analysis and pulsed-field gel electrophoresis for typing of Vibrio vulnificus and their correspondence with multilocus sequence typing data. Appl Environ Microbiol 73: 7494-500. 31. Gordon, K. V., M. C. Vickery, A. DePaola, C. Staley, and V. J. Harwood. 2008. Real-time PCR assays for quan tification and differentiation of Vibrio vulnificus strains in oysters and wate r. Appl Environ Microbiol 74: 1704-9. 32. Gray, L. D., and A. S. Kreger. 1986. Detection of antiVibrio vulnificus cytolysin antibodies in sera from mice and a human surviving V. vulnificus disease. Infect Immun 51: 964-5. 33. Gray, L. D., and A. S. Kreger. 1989. Detection of Vibrio vulnificus cytolysin in V. vulnificus -infected mice. Toxicon 27: 439-64. 34. Gray, L. D., and A. S. Kreger. 1985. Identification of Vibrio vulnificus by indirect immunofluorescence. Diagn Microbiol Infect Dis 3: 461-8.

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25 35. Gray, L. D., and A. S. Kreger. 1987. Mouse skin damage caused by cytolysin from Vibrio vulnificus and by V. vulnificus infection. J Infect Dis 155: 236-41. 36. Gray, L. D., and A. S. Kreger. 1985. Purification and characterization of an extracellular cytolysin produced by Vibrio vulnificus. Infect Immun 48: 62-72. 37. Gulig, P. A. 1993. Use of isogenic mutants to stu dy bacterial virulence factors. J Microbiol Meth 18: 275-287. 38. Gulig, P. A., K. L. Bourdage, and A. M. Starks. 2005. Molecular Pathogenesis of Vibrio vulnificus J Microbiol 43 Spec No: 118-131. 39. Haq, S. M., and H. H. Dayal. 2005. Chronic liver disease and consumption of raw oysters: a potentially lethal combination--a review of Vibrio vulnificus septicemia. Am J Gastroenterol 100: 1195-9. 40. Heid, C. A., J. Stevens, K. J. Livak, and P. M. Williams. 1996. Real time quantitative PCR. Genome Res 6: 986-94. 41. Helms, S. D., J. D. Oliver, and J. C. Travis. 1984. Role of heme compounds and haptoglobin in Vibrio vulnificus pathogenicity. Infect Immun 45: 345-9. 42. Higgins, C. F., G. F. Ames, W. M. Ba rnes, J. M. Clement, and M. Hofnung. 1982. A novel intercistronic regulatory el ement of prokaryotic operons. Nature 298: 760-2. 43. Higuchi, R., C. Fockler, G. Dollinger, and R. Watson. 1993. Kinetic PCR analysis: real-time monitoring of DNA am plification reactions. Biotechnology (N Y) 11: 1026-30. 44. Hlady, W. G., and K. C. Klontz. 1996. The epidemiology of Vibrio infections in Florida, 1981-1993. J Infect Dis 173: 1176-83. 45. Holbein, B. E., K. W. Jericho, and G. C. Likes. 1979. Neisseria meningitidis infection in mice: influence of iron, vari ations in virulen ce among strains, and pathology. Infect Immun 24: 545-51. 46. Hulton, C. S., C. F. Higgins, and P. M. Sharp. 1991. ERIC sequences: a novel family of repetitive elements in the genomes of Escherichia coli Salmonella typhimurium and other enterobacteria. Mol Microbiol 5: 825-34. 47. Jackson, J. K., R. L. Murphree, and M. L. Tamplin. 1997. Evidence that mortality from Vibrio vulnificus infection results from single strains among heterogeneous populations in shellfish. J Clin Microbiol 35: 2098-2101. 48. Johnson, D. E., and F. M. Calia. 1981. Hemolytic reaction of clinical and environmental strains of Vibrio vulnificus J Clin Microbiol 14: 457-9.

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26 49. Johnson, L. K., M. B. Brown, E. A. Carruthers, J. A. Ferguson, P. E. Dombek, and M. J. Sadowsky. 2004. Sample size, library composition, and genotypic diversity among natural populations of Escherichia coli from different animals influence accuracy of determin ing sources of fecal pollution. Appl Environ Microbiol 70: 4478-85. 50. Jones, M. K., E. Warner, and J. D. Oliver. 2008. Survival of and in situ gene expression by Vibrio vulnificus at varying salinities in estuarine environments. Appl Environ Microbiol 74: 182-7. 51. Kaper, J. B., A. Fasano, and M. Trucksis. 1994. Toxins of Vibrio cholerae, p. 145-176. In K. I. Wachsmuth, P. A. Blak e, and O. Olsvik (ed.), In: Vibrio cholerae and Cholera: Molecular to Global Perspectives. American Society for Microbiology, Washington, DC. 52. Kashimoto, T., S. Ueno, M. Hanajima, H. Hayashi, Y. Akeda, S. Miyoshi, T. Hongo, T. Honda, and N. Susa. 2003. Vibrio vulnificus induces macrophage apoptosis in vitro and in vivo. Infect Immun 71: 533-5. 53. Kaysner, C. A., C. Abeyta, Jr., M. M. Wekell, A. DePaola, Jr., R. F. Stott, and J. M. Leitch. 1987. Virulent strains of Vibrio vulnificus isolated from estuaries of the United States West Coast. Appl Environ Microbiol 53: 1349-51. 54. Kaysner, C. A., and A. DePaola. 2004. Vibrio cholerae V. parahaemolyticus V. vulnificus, and Other Vibrio spp., In Bacteriological Analytical Manual Online, 8th ed. Revision A, 1998. Chapter 9. Subs tantially rewritten and revised May 2004. http://www.cfsan.fda.gov/~ebam/bam-9.html 55. Kim, H. R., H. W. Rho, M. H. Jeong, J. W. Park, J. S. Kim, B. H. Park, U. H. Kim, and S. D. Park. 1993. Hemolytic mechanism of cytolysin produced from V. vulnificus. Life Sci 53: 571-7. 56. Kim, M. S., and H. D. Jeong. 2001. Development of 16S rRNA targeted PCR methods for the detectio n and differentiation of Vibrio vulnificus in marine environments. Aquaculture 193: 199-211. 57. Kim, Y. R., S. E. Lee, C. M. Kim, S. Y. Kim, E. K. Shin, D. H. Shin, S. S. Chung, H. E. Choy, A. Progulske-Fox, J. D. Hillman, M. Handfield, and J. H. Rhee. 2003. Characterization and pa thogenic significance of Vibrio vulnificus antigens preferentially expressed in septicemic patients. Infect Immun 71: 546171. 58. Kim, Y. R., and J. H. Rhee. 2003. Flagellar basal body flg operon as a virulence determinant of Vibrio vulnificus Biochem Biophys Res Commun 304: 405-410.

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27 59. Koeuth, T., J. Versalovic, and J. R. Lupski. 1995. Differential subsequence conservation of inte rspersed repetitive Streptococcus pneumoniae BOX elements in diverse bacter ia. Genome Res 5: 408-18. 60. Kook, H., S. E. Lee, Y. H. Ba ik, S. S. Chung, and J. H. Rhee. 1996. Vibrio vulnificus hemolysin dilates rat thoracic ao rta by activating guanylate cyclase. Life Sci 59: PL41-7. 61. Kreger, A., and D. Lockwood. 1981. Detection of ex tracellular toxin(s) produced by Vibrio vulnificus Infect Immun 33: 583-90. 62. Kwon, K. B., J. Y. Yang, D. G. Ryu, H. W. Rho, J. S. Kim, J. W. Park, H. R. Kim, and B. H. Park. 2001. Vibrio vulnificus cytolysin induces superoxide anion-initiated apoptotic signaling pathway in human ECV304 cells. J Biol Chem 276: 47518-23. 63. Lee, J. H., J. B. Rho, K. J. Park, C. B. Kim Y. S. Han, S. H. Choi, K. H. Lee, and S. J. Park. 2004. Role of flagellum and motility in pathogenesis of Vibrio vulnificus. Infect Immun 72: 4905-10. 64. Lee, S. E., S. Y. Kim, B. C. Jeong, Y. R. Kim, S. J. Bae, O. S. Ahn, J. J. Lee, H. C. Song, J. M. Kim, H. E. Choy, S. S. Chung, M. N. Kweon, and J. H. Rhee. 2006. A bacterial flagellin, Vibrio vulnificus FlaB, has a strong mucosal adjuvant activity to induce prot ective immunity. Infect Immun 74: 694-702. 65. Lee, S. E., S. Y. Kim, S. J. Kim, H. S. Kim, J. H. Shin, S. H. Choi, S. S. Chung, and J. H. Rhee. 1998. Direct identification of Vibrio vulnificus in clinical specimens by nested PCR. J Clin Microbiol 36: 2887-2892. 66. Lee, S. E., P. Y. Ryu, S. Y. Kim, Y. R. Kim, J. T. Koh, O. J. Kim, S. S. Chung, H. E. Choy, and J. H. Rhee. 2004. Production of Vibrio vulnificus hemolysin in vivo and its pathogeni c significance. Biochem Biophys Res Commun 324: 86-91. 67. Levin, R. E. 2006. Vibrio parahaemolyticus a notably lethal human pathogen derived from seafood: A review of its pa thogenicity, characteristics, subspecies characterization, and molecular met hods of detection. Food Biotechnology 20. 68. Linkous, D. A., and J. D. Oliver. 1999. Pathogenesis of Vibrio vulnificus FEMS Microbiol Lett 174: 207-214. 69. Litwin, C. M., T. W. Rayback, and J. Skinner. 1996. Role of catechol siderophore synthesis in Vibrio vulnificus virulence. Infect Immun 64: 2834-8. 70. Maiden, M. C., J. A. Bygraves, E. Feil, G. Morelli, J. E. Russell, R. Urwin, Q. Zhang, J. Zhou, K. Zurth, D. A. Caugant, I. M. Feavers, M. Achtman, and B. G. Spratt. 1998. Multilocus sequenc e typing: a portable approach to the

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28 identification of clones within populations of pathogenic microorganisms. Proc Natl Acad Sci U S A 95: 3140-5. 71. Martin, B., O. Humbert, M. Camara, E. Guenzi, J. Walker, T. Mitchell, P. Andrew, M. Prudhomme, G. Alloing, R. Hakenbeck, and et al. 1992. A highly conserved repeated DNA element located in the chromosome of Streptococcus pneumoniae Nucleic Acids Res 20: 3479-83. 72. Motes, M. L., A. DePaola, D. W. Cook, J. E. Veazey, J. C. Hunsucker, W. E. Garthright, R. J. Blodgett, and S. J. Chirtel. 1998. Influence of water temperature and salinity on Vibrio vulnificus in Northern Gulf and Atlantic Coast oysters ( Crassostrea virginica ). Appl Environ Microbiol 64: 1459-1465. 73. Nilsson, W. B., R. N. Paranjype, A. DePaola, and M. S. Strom. 2003. Sequence polymorphism of the 16S rRNA gene of Vibrio vulnificus is a possible indicator of strain virulence. J Clin Microbiol 41: 442-446. 74. Nishibuchi, M., A. Fasano, R. G. Russell, and J. B. Kaper. 1992. Enterotoxigenicity of Vibrio parahaemolyticus with and without genes encoding thermostable direct hemolysin. Infect Immun 60: 3539-45. 75. Nishibuchi, M., and J. B. Kaper. 1985. Nucleotide sequence of the thermostable direct hemolysin gene of Vibrio parahaemolyticus. J Bacteriol 162: 558-64. 76. Nishibuchi, M., T. Taniguchi, T. Mi sawa, V. Khaeomanee-Iam, T. Honda, and T. Miwatani. 1989. Cloning and nucleotide sequence of the gene ( trh ) encoding the hemolysin related to the thermostable direct hemolysin of Vibrio parahaemolyticus Infect Immun 57: 2691-7. 77. Nordstrom, J. L., M. C. Vickery, G. M. Blackstone, S. L. Murray, and A. DePaola. 2007. Development of a multiplex realtime PCR assay with an internal amplification control for the de tection of tota l and pathogenic Vibrio parahaemolyticus bacteria in oysters. A ppl Environ Microbiol 73: 5840-7. 78. Okujo, N., T. Akiyama, S. Miyoshi, S. Shinoda, and S. Yamamoto. 1996. Involvement of vulnibactin and exocellular protease in utilization of transferrinand lactoferrin-bound iron by Vibrio vulnificus Microbiol Immunol 40: 595-8. 79. Okujo, N., M. Saito, S. Yamamoto, T. Yoshida, S. Miyoshi, and S. Shinoda. 1994. Structure of vulnibactin, a new pol yamine-containing siderophore from Vibrio vulnificus Biometals 7: 109-16. 80. Panicker, G., D. R. Call, M. J. Krug, and A. K. Bej. 2004. Detection of pathogenic Vibrio spp. in shellfish by using multiplex PCR and DNA microarrays. Appl Environ Microbiol 70: 7436-44.

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29 81. Panicker, G., M. C. Vickery, and A. K. Bej. 2007. Erratum: Multiplex PCR detection of clinical and environmental strains of Vibrio vulnificus in shellfish. Can J Microbiol 53: 671. 82. Panicker, G., M. C. Vickery, and A. K. Bej. 2004. Multiplex PCR detection of clinical and envir onmental strains of Vibrio vulnificus in shellfish. Can J Microbiol 50: 911-22. 83. Paranjpye, R. N., J. C. Lara, J. C. Pepe, C. M. Pepe, and M. S. Strom. 1998. The type IV leader peptid ase/N-methyltransferase of Vibrio vulnificus controls factors required for adherence to HEp-2 cells and virulence in iron-overloaded mice. Infect Immun 66: 5659-68. 84. Reidl, J., and K. E. Klose. 2002. Vibrio cholerae and cholera: out of the water and into the host. FEMS Microbiol Rev 26: 125-39. 85. Rosche, T. M., Y. Yano, and J. D. Oliver. 2005. A rapid and simple PCR analysis indicates ther e are two subgroups of Vibrio vulnificus which correlate with clinical or environmenta l isolation. Microbiol Immunol 49: 381-389. 86. Saiki, R. K., S. Scharf, F. Faloona, K. B. Mullis, G. T. Horn, H. A. Erlich, and N. Arnheim. 1985. Enzymatic amplification of beta-globin genomic sequences and restriction site analysis for diagnosis of sickle cell anemia. Science 230: 1350-4. 87. Shapiro, R. L., S. Altekruse, L. Hu twagner, R. Bishop, R. Hammond, S. Wilson, B. Ray, S. Thompson, R. V. Tauxe, and P. M. Griffin. 1998. The role of Gulf Coast oysters harves ted in warmer months in Vibrio vulnificus infections in the United States, 1988-1996. Vibrio Working Group. J Infect Dis 178: 752-9. 88. Sharples, G. J., and R. G. Lloyd. 1990. A novel repeated DNA sequence located in the intergenic regions of bacterial chromosomes. Nucleic Acids Res 18: 6503-8. 89. Shinoda, S., S. Miyoshi, H. Yamanaka, and N. Miyoshi-Nakahara. 1985. Some properties of Vibrio vulnificus hemolysin. Microbiol Immunol 29: 583-90. 90. Simpson, L. M., and J. D. Oliver. 1983. Siderophore production by Vibrio vulnificus. Infect Immun 41: 644-9. 91. Simpson, L. M., V. K. White, S. F. Zane, and J. D. Oliver. 1987. Correlation between virulence and colony morphology in Vibrio vulnificus Infect Immun 55: 269-272. 92. Starks, A. M., K. L. Bourdage, P. C. Thiaville, and P. A. Gulig. 2006. Use of a marker plasmid to examine differential rate s of growth and death between clinical and environmental strains of Vibrio vulnificus in experimentally infected mice. Mol Microbiol 61: 310-23.

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30 93. Starks, A. M., T. R. Schoeb, M. L. Tamplin, S. Parveen, T. J. Doyle, P. E. Bomeisl, G. M. Escudero, and P. A. Gulig. 2000. Pathogenesis of infection by clinical and envir onmental strains of Vibrio vulnificus in iron-dextran-treated mice. Infect Immun 68: 5785-93. 94. Stelma, G. N., Jr., A. L. Reyes, J. T. Peeler, C. H. Johnson, and P. L. Spaulding. 1992. Virulence characteristics of c linical and environmental isolates of Vibrio vulnificus Appl Environ Microbiol 58: 2776-82. 95. Strom, M. S., and R. N. Paranjpye. 2000. Epidemiology and pathogenesis of Vibrio vulnificus Microbes Infect 2: 177-88. 96. Tacket, C. O., F. Brenner, and P. A. Blake. 1984. Clinical features and an epidemiological study of Vibrio vulnificus infections. J Infect Dis 149: 558-61. 97. Tamplin, M. L., and G. M. Capers. 1992. Persistence of Vibrio vulnificus in tissues of Gulf Coast oysters, Crassostrea virginica exposed to seawater disinfected with UV light. Appl Environ Microbiol 58: 1506-10. 98. Tamplin, M. L., J. K. Jackson, C. Buchries er, R. L. Murphree, K. M. Portier, V. Gangar, L. G. Miller, and C. W. Kaspar. 1996. Pulsed-field gel electrophoresis and ribotype profiles of clinical and environmental Vibrio vulnificus isolates. Appl Environ Microbiol 62: 3572-3580. 99. Tison, D. L., M. Nishibuchi, J. D. Greenwood, and R. J. Seidler. 1982. Vibrio vulnificus biogroup 2: new biogroup pathogenic fo r eels. Appl Environ Microbiol 44: 640-646. 100. Tyagi, S., and F. R. Kramer. 1996. Molecular beacons: probes that fluoresce upon hybridization. Nat Biotechnol 14: 303-8. 101. Versalovic, J., T. Koeuth, and J. R. Lupski. 1991. Distribution of repetitive DNA sequences in eubacteria and applic ation to fingerprint ing of bacterial genomes. Nucleic Acids Res 19: 6823-31. 102. Vickery, M. C., W. B. Nilsson, M. S. Strom, J. L. Nordstrom, and A. DePaola. 2007. A real-time PCR assay for the rapid determination of 16S rRNA genotype in Vibrio vulnificus J Microbiol Methods 68: 376-384. 103. Warner, E., and J. D. Oliver. 2008. Population structures of two genotypes of Vibrio vulnificus in oysters ( Crassostrea virginica) and seawater. Appl Environ Microbiol 74: 80-5. 104. Warner, J. M., and J. D. Oliver. 1998. Randomly amplified polymorphic DNA analysis of starved and viable but nonculturable Vibrio vulnificus cells. Appl Environ Microbiol 64: 3025-8.

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31 105. Weinberg, E. D. 1978. Iron and infection. Microbiol Rev 42: 45-66. 106. Weinberg, E. D. 2000. Microbial pathogens with im paired ability to acquire host iron. Biometals 13: 85-9. 107. Wright, A. C., R. T. Hill, J. A. Johnson, M. C. Roghman, R. R. Colwell, and J. G. Morris, Jr. 1996. Distribution of Vibrio vulnificus in the Chesapeake Bay. Appl Environ Microbiol 62: 717-724. 108. Wright, A. C., G. A. Miceli, W. L. Landry, J. B. Christy, W. D. Watkins, and J. G. Morris, Jr. 1993. Rapid identification of Vibrio vulnificus on nonselective media with an alkaline phosphatase-labeled oligonucle otide probe. Appl Environ Microbiol 59: 541-546. 109. Wright, A. C., and J. G. Morris, Jr. 1991. The extracellular cytolysin of Vibrio vulnificus : inactivation and relationship to virulence in mice. Infect Immun 59: 192-197. 110. Wright, A. C., J. G. Morris, Jr., D. R. Maneval, Jr., K. Richardson, and J. B. Kaper. 1985. Cloning of the cytotoxin-hemolysin gene of Vibrio vulnificus Infect Immun 50: 922-924. 111. Wright, A. C., L. M. Simpson, and J. D. Oliver. 1981. Role of iron in the pathogenesis of Vibrio vulnificus infections. Infect Immun 34: 503-7. 112. Wright, A. C., L. M. Simpson, J. D. Oliver, and J. G. Morris, Jr. 1990. Phenotypic evaluation of acapsular transposon mutants of Vibrio vulnificus Infect Immun 58: 1769-1773. 113. Yamamoto, K., A. C. Wright, J. B. Kaper, and J. G. Morris, Jr. 1990. The cytolysin gene of Vibrio vulnificus : sequence and relationship to the Vibrio cholerae E1 Tor hemolysin gene. Infect Immun 58: 2706-9. 114. Yoshida, S., M. Ogawa, and Y. Mizuguchi. 1985. Relation of capsular materials and colony opacity to virulence of Vibrio vulnificus. Infect Immun 47: 446-451. 115. Zuppardo, A. B., and R. J. Siebeling. 1998. An epimerase gene essential for capsule synthesis in Vibrio vulnificus Infect Immun 66: 2601-6.

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32 REAL-TIME PCR ASSAYS FOR QUANT IFICATION AND DIFFERENTIATION OF V. VULNIFICUS STRAINS IN OYSTERS AND WATER Katrina V. Gordon1, Michael C. Vickery2, Angelo DePaola3 Christopher Staley1 and Valerie J. Harwood1* 1Department of Biology, University of South Florida, Tampa, Florida, 33620, 2Life Sciences Division, Cepheid, Inc., Sunnyvale, CA 94089 and 3Gulf Coast Seafood Laboratory, U.S. Food and Drug Admini stration, Dauphin Island, Alabama, 36528 Running title: Real-time PCR for V. vulnificus typing Key words: Vibrio vulnificus real-time PCR, food safet y, water quality, shellfish, genotype *Corresponding author. Mailing address: Department of Biology, Univ ersity of South Florida, 4202 E Fowler Ave, SCA 110, Tampa, FL 33620. Phone: (813) 974-1524. Fax: (813) 974-3263. E-mail: vharwood@cas.usf.edu (Published in Applied and Environm ental Microbiology Vol. 74 (6):1704-1709)

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33 Abstract Vibrio vulnificus is an autochthonous estuarine ba cterium and a pathogen that is frequently transmitted via raw shellfish. Se pticemia can occur within 24 h; however, isolation and confirmation from water and oys ters require days. Real-time PCR assays were developed to detect and differentiate two 16S rRNA variants, type A and B, which were previously associated with envir onmental sources and clinical fatalities, respectively. Both assays could detect 102-103 V. vulnificus total cells in seeded estuarine water and in oyster homogenates. PCR assays on 11 reference V. vulnificus strains and 22 nontarget species gave expected results (type A or B for V. vulnificus and negative for nontarget species). The relati onship between cell number and cycle threshold for the assays was linear (R2 > 0.93). The type A:B ratio of Florida clinical isolates was compared to that of isolates from oysters harvested in Florida wa ters. This ratio was 19:17 in clinical isolates and 5:8 (n=26) in oysters harvested from restricted sites with poor water quality, but was 10:1 (n=22) in oysters from permitted sites with good water quality. A substantial percentage of isolates from oysters (19.4%) were type AB (both primer sets amplified), but no isolates from overlying waters were type AB. The real-time PCR assays were sensitive, specific, and qua ntitative in water samples and could also differentiate the strains in oysters without requiri ng isolation of V. vulnificus, and may therefore be useful for rapid detection of the pathogen in shellfish and water, as well as further investigation of its population dynamics.

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34 Introduction Vibrio vulnificus is a gram-negative bacterium that is autochthonous to warm estuarine waters and is frequently isolated from shellfish harvested in the Gulf of Mexico. V. vulnificus infections have been noted as the le ading cause of food-related mortality in Florida (12). In the USA, nearly all foodborne infections result from the consumption of oysters collected from the Gulf of Mexico (5). Immunocompromis ed individuals and those with diseases causing increased iron le vels in the body, such as liver disease or hemochromatosis, are at relatively high risk for development of primary septicemia, with a mortality rate of around 50% (6). There is al so a risk of acute gastroenteritis due to the consumption of raw or undercooked seafood such as oysters (9). Infec tions can also occur due to trauma associated with handling contaminated seafood or the contact of open wounds with water containing V. vulnificus Wound infections can become fatal or so severe that amputation is necessary to st op the spread of infection (17). Although the health of the host is a factor, it is not an absolute determinant of infection or clinical outcome (26). Regulations for water quality in shellf ish harvesting areas in Florida rely on testing for indicator organisms, i.e. fecal coliforms (http://www.floridaaqua culture.com/SEAS/SEAS_intro.htm). For the most part, indicator bacteria have not been shown to corr elate with the presence of pathogenic Vibrio spp. (13, 22, 27). At present, proposed risk assessment models rely on testing for the total V. vulnificus population (10). Currently accepted methods for isolation of V. vulnificus from seafood require plating on one of several se lective-differential media (11, 14), followed by confirmation by biochemical or molecula r tests (4, 11, 14, 15, 32). These methods

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35 require at least 24 h for completion, and do not take into account the potential for variation in strain virulence. Efforts to identify virulence factors for V. vulnificus have met with varying success. The capsular polysaccharid e is present in nearly all V. vulnificus strains at the time of isolation, although translucent varian ts may arise during laboratory culture that are avirulent (28, 34). One study (33) showed th at inactivation of the cytolysin gene did not affect the LD50 of virulent V. vulnificus strains. Sequence polymorphism of the 16S rRNA gene (3) was used to develop a restri ction fragment polymorphism (RFLP) method to differentiate V. vulnificus strains (19). Typing of V. vulnificus isolates grouped environmental isolates in RFLP type A (94%), while the majority of those from clinical cases were RFLP type B (76%), as were 94.4% of strains from clinical fatalities (19). In this study, we developed primers for use in a SYBR green based, real-time PCR assay to detect and diffe rentiate rRNA types A and B V. vulnificus without an isolation requirement. Development of a rapid, cost e ffective test for the relatively virulent strain(s) of this pathogen would aid in more accurate exposur e assessment and perhaps more appropriate allocation of resources to mitigate the risk. Methods Bacterial strains Vibrio vulnificus strains of clinical orig in (n=36) and non-target isolates (n=22) were obtained from th e culture collection of the Food and Drug Administration (FDA) Gulf Coast Seafood Laboratory, from Dr. Daniel Lim (University of South Florida, Tampa), from Dr. A. Cannons and Mr. R. Baker (Florida Department of

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36 Health, FDOH), from the Centers for Diseas e Control (CDC) and from the American Type Culture Collection (ATCC). The eleven reference strains used to establish agreement among the typing methods were clin ical or environmental strains obtained from the FDA (ten strains), and from ATCC (27562). Environmental strains (n=82) were collected from the Guana-Tolomato-Matanzas National Estuarine Research Reserve (GTMNERR) near St. Augustine on the east coast of Florida during four sample events, from Tampa Bay during two sample events, and from Apalachicola, FL during one sample event. Samples collected from St. Augustine and Tampa Bay were collected from non-permitted oyster harvesting areas when th e water temperature was generally above 24C. Oyster samples collected from Apalachicola were harvested by a commercial shell fishing company from permitted oyster harvesting areas in August, when water temperatures were also >24C. Oyster homoge nates were prepared from oysters diluted 1:10 (w/w) in alkaline peptone water (APW). All putative V. vulnificus isolates were confirmed by PCR using the vvhA gene (15). Specificity assays were conducted on 22 non-target bacterial species including closely related Vibrios ( V. alginolyticus ATCC 51160, V. cholerae El Tor, O1, ATCC 25780, V. parahaemolyticus ATCC 10290 and 49398 Aeromonas hydrophila ATCC 7965 and 14715 as well as members of the co liform groups commonly found in marine and estuarine waters ( Citrobacter freundii ATCC 8090, Enterobacter agglomerans ATCC 27981, Enterococcus avium Enterococcus casseliflavus Enterococcus faecalis Enterococcus faecium Enterococcus gallinarum Enterococcus mundtii Klebsiella ozaenae ATCC 29019, Klebsiella pneumoniae ATCC 13883, Pseudomonas aeruginosa

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37 ATCC 15442, Serratia marcescens ATCC 14756, Weisella confusa ATCC14434, Yersinia enterocolitica ATCC 9610). Preparation of pure cultures For specificity, sensitivity and typing assays, Vibrio spp. and other isolates were grown in 25 ml brain heart infusion broth (BHI) supplemented with 0.5% NaCl for 1824 h at 25C with shaking. For sensitivity assays, V. vulnificus ATCC 33814 and vPvMH1003-12 were concentrated by centrifugation at 14,500 g for 10 min. The supernatant was removed and the pellet was washed twice with 2.0 ml of 0.85% NaCl before being re suspended in 4.0 ml of 0.85% NaCl. Serial dilutions were then made in 0.85% NaCl, or in an alternative matrix as noted. Alternative matrices included filter-sterilized Instant Ocean (20) (Aquarium Systems Inc., Mentor, OH), or unsterilized water collected from Tampa Bay, FL (pH 7.96, salinity 28 and temperature of 21C at time of collection). The concentration of cells was determined using total direct microscopic counts (see be low), and consistent volumes of successive dilutions were then used as the template in real-time PCR assays. DNA was extracted only for the specificity and typing assays using a DNeasy Tissue Kit (Qiagen) according to the manufacturers instructi ons, and quantified with a Beckman DU640 spectrophotometer (Beckman C oulter) using a 2:7 dilution in 10 mM Tris-HCl (pH 8.5). Total cell counts Dilution series were made in 0.85% NaCl (or appropriate medium) as above for direct fluorescence microscopic counts. DAPI (4,6-diamidine-2phenylindole) staining was performed essentiall y as previously described (23) using 1l of DAPI (1mg ml-1) to stain 1 ml samples. Each dilution was filtered through a 0.2m GTBP Isopore membrane filter (Millipor e, Burlington, MA) and observed under a Nikon Diaphot inverted fluorescen ce microscope. The number of cells in a 20 square area

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38 (0.002 mm2) of the ocular grid was recorded fo r 5 different fields of view, and cell concentrations were calculated. Primer development Sequences for two 16S rRNA variants of V. vulnificus identified by Aznar et al (3) were obtained from GenBank (accession numbers X67333 and X76334). The two sequences were aligned using GeneDoc (18) to identify the positions of the 17 base pair differences. Forward primer sequences were adapted from probes previously designed by Michael Vickery et al. (30); the sequence of the type A forward primer was VvAF1 5 -CAT GAT AGC TTC GGC TCA A-3 and that of the B forward primer was VvBF1 5 -GCC TAC GGG CCA AAG AGG-3 The positions of the reverse primers (VvAR1 5 CAG CAC TCC TTC CAC CAT CAC-3 and VvBR1 5 GTC GCC TCT GCG TCC AC-3 ) were chosen with the aid of PrimerQuest (24) to allow easily differentiable PCR products (type A=245 bp and type B=841 bp) while capitalizing on as many of the available base pair variations as possible. Initially, the specificity of the primers was assessed us ing BLAST, National Center for Biotechnology Information (2) which showed 100% similarity for each primer set only with V. vulnificus sequences. PCR conditions were optimized and then adapted for use with SYBR Green (Roche Diagnostics, Germany) in a real -time PCR assay with the LightCycler 2.0 (Roche). Real-time PCR protocol Real-time PCR assays of 20 l contained 2 mM MgCl2, 0.5 M of each primer, 2 l of LightCycler FastStart DNA Master SYBR Green I reaction mix (Roche Diagnostics, Germany), a nd PCR template (whole cells or extracted DNA). When purified DNA was used as the template (for specificity and typing analysis), 2 l of template solution containing 420 ng of DNA was added to each

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39 reaction. Whole cells were used for sensitivity analysis. Po sitive and negative controls were included for each set of reactions: for type A primers, an environmental V. vulnificus isolate (vPvMH1003-12) was used and for type B primers, V. vulnificus ATCC 33814 was used. An initial 10 min. denaturation step at 95 C was followed by a two-step amplification program of 35 cycles of 95 C for 15s and 74C for 35s. Triplicate reactions were performed to confirm reproducibility of results. PCR products for both primer sets were also sequenced using GenomeLab DTCS Quick Start Kit (Beckman Coulter, Fullerton CA.) to ensure the correct amplicons were produced. RFLP confirmation Restriction fragment length polymorphism (RFLP) was performed on PCR products from eleven reference V. vulnificus strains using Alu I and Hae III as previously described (19) in or der to validate the PCR typing method. Sensitivity. V. vulnificus concentrations were determined by direct microscopic counts. Cells were diluted in 0.85% NaCl in order to add from 106 cells to 1 cell to each PCR reaction. Reactions were also carried out usi ng a mixture of an equal number of cells from V. vulnificus ATCC 33814 and vPvMH1003-12. To te st primer sensitivity in more natural matrices, dilution series were al so made using filter sterilized 20 Instant Ocean (Aquarium Systems Inc., Mentor, OH), and estuarine water (unsterilized water from Tampa Bay) as above. Type A and type B control V. vulnificus cells were inoculated into 300 ml of water collected from Tampa Bay (salinity ~28 ) to a final concen tration of ~160 cells/ 100 ml. The estuarine wa ter was assayed for V. vulnificus prior to the experiment and did not contain detectable levels of the bacterium. The seeded water was filtered through a 0.45 m pore size nitrocellulose filter (Osmonics Inc, Westborough, MA). Genomic

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40 DNA was extracted from the filter usi ng the UltraClean Soil DNA Kit (Mo Bio Laboratories Inc., Solana Beach, CA). The final elution volume was 50 l (6000-fold nominal concentration facto r). Two microliters of the extracted DNA was used as template in triplicate real-time PCR assays as above. The sensitivity of the assays in homogen ized oyster tissue was tested by seeding 2 l of oyster homogenate (see be low), which had not previous ly shown any amplification with the primer sets, with 1 to 107 cells of type A or type B control V. vulnificus The seeded homogenate was then direc tly analyzed by real-time PCR. PCR detection of native V. vuln ificus in oyster homogenates Tissue from a single oyster collected from Tampa Bay was diluted 1: 10 (w/w) in alkaline peptone water (14). Homogenate was made by blending at high sp eed for 90s. A 25 ml a liquot was pipetted into two centrifuge tubes for enrichment at 37C for 4 and 24 h. Enrichments and oyster homogenate were immediately cultured (see be low) and aliquots were stored at -20C for later testing by the real-time PCR. To determine the concentrations of culturable V. vulnificus naturally present in oyster homogenates, V. vulnificus was isolated from initial homogenates and dilutions of enriched homogenates by spreading 0.1 ml onto V. vulnificus agar (VVA) (14). VVA plates were incubated overnight at 37C in order to determine the presence of type A and/or type B V. vulnificus in each oyster. Cellobiose fermenting colonies from VVA plates were transferred indi vidually to 96 well microtiter plates containing 180 l T1N1 agar and stored at room temp erature for further analysis. Th e same set of VVA plates was

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41 then probed according to Wright et al. (32) w ith hybridization at 56C. The colonies that gave positive reactions with the probe were then typed by real-time PCR. The original oyster homogenate and enriched homogenates (4 h and 24 h enrichment) were prepared for PCR assays by diluting 1:10 (vol/vol) in 0.85% NaCl. Two milliliters of each diluted homogenate wa s boiled for 15 min. to lyse cells and centrifuged at 16,000 x g to pellet cellular debr is. Two microliters of the supernatant was the template for each PCR reaction. To monito r for possible inhibition of amplification from oyster homogenates, positive control reactions were made which included 10-50 ng of V. vulnificus DNA (type A and type B). Statistical analysis Linear regression of cycle threshold (Ct value) vs. seeded cell numbers (enumerated by direct microscope count) was conducted in Microsoft Excel. A contingency table and Chi-squared test (Gra phPad InStat v. 3.00) was used to compare the A:B:AB ratio of V. vulnificus isolates. Results Both primer sets performed as expected, as each amplified the 16S rDNA from the corresponding type of V. vulnificus and no product was produced when the template was the opposite V. vulnificus type. Agarose gel electr ophoresis of PCR products indicated that amplicons were of the expected sizes (285 bp for type A, and 841 bp for type B) and had average melti ng temperatures of 89.0.3 C for products of type A and type B primer sets. BLAST (2) analysis showed that the DNA sequence of the PCR products produced by each primer set was most closely related to its respective V.

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42 vulnificus strain in GenBank. Multiplex assays using the A and B primers in the same reaction were not extensively explored due to the possibility of amplification of V. parahaemolyticus DNA in a multiplex format. The specificity of each primer set was tested against 22 non-target organisms which included closely related Vibrio spp. (see Methods section) as well as clinical and environmental V. vulnificus strains (Tables 1 and 2). No amplification was observed with any of the non-target bacterial isolates tested. DNA from each V. vulnificus isolate was amplified, confirming it as type A, type B or in some cases, type AB (Tables 1 and 2); the latter denotes positive results with both the A and B primer sets in separate reactions.

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43 Table 1. V. vulnificus strains, origin, and type. Type A isolates obtained from clinical samples in Florida are bolded. Superscripts (see below) designate source of infection when known. PCR PCR Strain Source Result Strain Source Result ATCC 27562 c Clinical, Fl. A ATCC 33814 Clinical, Fl. B VV 9031-96 a,c FDA, Clinical,Fl A VV 9053-96 a,c FDA, Clinical, TX B 4933b FDOH, Clinical, FL A VV 9067-96 a,c FDA, Clinical, TX B 4939a FDOH, Clinical, FL A 6130a FDOH, Clinical, FL B 5011 FDOH, Clinical, FL A 6283b FDOH, Clinical, FL B 5192b FDOH, Clinical, FL A 6434b FDOH, Clinical, FL B 6288a FDOH, Clinical, FL A 4263 FDOH, Clinical, FL B 6325b FDOH, Clinical, FL A 4265a FDOH, Clinical, FL B 6689 FDOH, Clinical, FL A 4350 FDOH, Clinical, FL B CBD 113 FDOH, Clinical, FL A 4351 FDOH, Clinical, FL B 1497-82 CDC, Clinical, Romania A 4352 FDOH, Clinical, FL B 1498-82 CDC, Clinical, Romania A 5274b FDOH, Clinical, FL B 2415-01 CDC, Clinical, TN A T&CH 83104-MT#1 FDOH, Clinical, FL B 2430-01 CDC, Clinical, FL A T&CH 83104-MT#2 FDOH, Clinical, FL B 2432-02 CDC, Clinical, LA A 2428-01 CDC, Clinical, UT B 2438-02 CDC, Clinical, CO A 2431-01 CDC, Clinical, NYC B 2448-03 CDC, Clinical, VA A 2450-06 CDC, Clinical, HI B 2428-06 CDC, Clinical, LA A 2432-06 CDC, Clinical, LA A a Infection through ingestion of contaminated oysters. b Wound infection. c Included in reference strain set

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44 Table 2. Genotypes of V. vulnificus isolates from oysters, wa ter and clinical sources. Type Isolate Source A B AB Total Environmental oysters water oysters water oysters water oysters water Tampa Bay 8 5 13 4 5 0 26 9 St. Augustine 2 4 3 2 0 0 5 6 Apalachicola 20 0 2 0 8 0 30 0 Previously typeda 5 0 1 0 0 0 6 0 Total 35 9 19 6 13 0 67 15 Clinical 19 17 0 36 Total 63 42 13 118 aReferences 19 and 30 RFLP confirmation Of the eleven reference strains that were also typed by RFLP using both restriction enzymes, nine strains re turned the same type with all methods. For two isolates, the Hae III results agreed with the re al-time PCR type however the Alu I pattern was a mixture of the ba nding patterns expected for type A and type B indicating a type AB result which has been seen previously (30). Sensitivity. Assay sensitivity was determined as the minimum number of seeded V. vulnificus cells required for amplification in 3 replicate PCR reactions. Cell numbers in pure cultures were assessed by direct mi croscopic counts, providing a more accurate and sensitive estimate of tota l cell concentrations than woul d culturable counts. In 0.85% NaCl with and without an equal number of non-target type B cells added, the

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45 VvAF1/VvAR1 primer set (type A) was able to quantify 102 to 106 cells of V. vulnificus vPvMH1003-12. In these same matrices/ treatm ents, primer set VvBR1/VvBF1 (type B) was consistently able to quantify 10 106 V. vulnificus ATCC 33814 cells per PCR assay. Linear regression showed an excellent correlation between cell number and Ct (cycle threshold), average R2=0.97 0.04 for both assays. The assay sensitivity was 103 cells for type A and 102 cells for type B primers se ts in 20 Instant Ocean (R2 0.93) or unsterilized estuarine water (R2 0.98). This demonstrates the potential usefulness of these assays in quantifying the two V. vulnificus types in estuarine waters. In our laboratory, the comparison of culturable to di rect microscopic cell counts for an 18-hour culture in BHI broth + 0.5% NaCl showed culturable counts were ~102 lower than direct counts (data not shown). In terms of cultura ble cells, this increases the apparent sensitivity in estuarine waters to 10 CFU pe r reaction for the type A assay and 1 CFU per reaction for the type B assay. To demonstrate that cell c oncentrations representative of levels found in natural environments could be detected by the PCR assays, V. vulnificus cells were seeded into 300 ml of estuarine water and were subseque ntly concentrated by membrane filtration and detected by real-time PCR. Amplification of samples concentrated from estuarine waters seeded with 160 cells100 ml-1 was observed in tripli cate reactions (each representing analysis of a samp le of ~12 ml before concentr ation) containing ~20 cells of either type A or type B V. vulnificus The Ct values obtained in this experiment were consistent with standard curves for both typing assays, which estimated ~17.4 and ~4 cells per reaction for type A and B respectively.

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46 In seeded oyster homogenates, the type A primer set was able to detect 103 to 106 cells per reaction while the t ype B primers could detect 102 to 106 cells per reaction. PCR detection of native V. vuln ificus in oyster homogenates Amplification from an unseeded oyster homogenate was achieved with the type B primer set after enrichment in APW for 24 h. Before enrichment the concentration of culturable V. vulnificus in the homogenate, which was derived fr om a single oyster, was 130 CFU g-1 oyster tissue, or 13 CFU ml-1 in the 1:10 homogenate. This en riched homogenate contained 1.2 107 CFU ml-1 V. vulnificus, or 2.4 104 CFU per PCR assay. In c ontrast, no amplification with type B primers was detected from th e homogenate without enrichment, or with enrichment for 4 h (which contained 2.5 103 CFU V. vulnificusml-1 enrichment). Furthermore, none of the reactions using th e type A primers yielded amplification. Typing of the naturally-occurring V. vulnificus strains isolated from the oyster homogenate yielded 100% type B (n = 9), which explains th e failure of amplification with type A primers. The lack of amplif ication by type B primers observed in the unenriched oyster homogenate and 4 hour-enric hed oyster homogenates was due to the relatively low number of naturally occurring V. vulnificus in this oyster. The oyster homogenate prior to enrichment contained 13 CFU ml-1 V. vulnificus, or 0.026 culturable cells per PCR reaction. After a 4-hour enrichment this increased to 5 culturable cells per PCR reaction. Typing of clinical and environmental isolates Thirty-six clinical isolates were typed in this study (Table 1). Fifty-three percent of these isolates were type A, and 47% were type B. Of the 32 clinical isolates not typed in previous st udies (19, 30), 17 (53%) were type A and 15 (47%) were type B. Ch i-squared analysis showed no significant

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47 difference between the relative frequency of the types and the distribution that would be expected by chance (P =0.8025), i.e. neither type compri sed a greater proportion of the clinical isolates. The route of infection for 13 of the clinical isolates not typed in previous studies was known. Six were the result of w ound infections and seven were from oyster consumption (Table 1). Seventy-six environmental is olates from water and oysters were typed that were not typed in previous studie s. All water samples (n=15) were from Tampa Bay or St. Augustine, where shellfishing is prohibited due to poor water quality in both areas. Sixty percent of the fifteen isolates from prohibite d shellfishing waters were type A, and 40% were type B (Table 2). Chi-squared analysis showed no significa nt difference in the observed frequency of isolation of each type from water and that expected by chance ( P =0.7144) Sixty-seven isolates from oysters were typed (Table 2). In oysters from Tampa and St. Augustine the majority of isolates were type B (the frequency of A:B:AB = 10:16:5; n = 31). To determine whether the freq uency of the types is olated from oysters was different in prohibited vs. permitted shellfishing waters, Chi-squared analysis was performed and the frequencies were found to differ significantly (Chi-square = 14.902, df =2, P =0.0006). Type B V. vulnificus were more frequently isolated from prohibited waters than from permitted wate rs (16:2). In contrast, type A V. vulnificus were more frequently isolated from oysters in permitted waters (prohibited:permitted = 10:20). Interestingly, type AB isolates were isolat ed at about the same frequency from oysters harvested from prohibited vs. permitted wate rs (5:8). Although type AB strains complicate the estimation of the relative frequency and numbers of V. vulnificus strains,

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48 they are in the minority in V. vulnificus populations [e.g. 21% of the isolates tested by Vickery et al. (30)]. Discussion The primers developed were able to amplify DNA from 100% of the V. vulnificus isolates, identifying them as type A, type B or type AB. RFLP analysis of 11 previously typed V. vulnificus isolates (19, 30) showed 82% agr eement with the real-time PCR assay types, determining the isolates to be type A, B or AB. The two discrepancies occurred when the Alu I enzyme of the RFLP assay sp ecified a type AB while the Hae III RFLP results and the PCR assays specified type A. Other studies have also documented the presence of type AB isolates (25, 30) due to amplification of genetic material from one isolate by primers/probes for both type A and type B. V. vulnificus YJ016 contains nine rRNA operons (8), which are all type B according to their sequences in GenBank. However, cloning and sequencing of 492 bp of the 16S rDNA from one type AB isolate showed the presence of type A and type B 16S rRNA genes (30). Heterogeneity between multiple 16S rRNA operon copies has also be en observed within the same strain of several related bacteria including V. parahaemolyticus and V. cholerae (1). Sensitivity. V. vulnificus could be detected at 160 cel ls/100 ml (1.6 cells/ml) in a water sample. Other studies have developed methods to detect V. vulnificus in water samples using real-time PCR targeting genes such as the cytolysin/hemolysin and toxR (4, 20, 21, 29, 31). The sensitivity of these methods ranged from the equivalent of 101 to 102 CFU ml-1 in seeded water samples (4, 21, 29) when using extracted DNA as the template. The sensitivity of our assay in wate r samples was greater than those previously

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49 reported. The assays are not as sensitive or rapid in oyster homogena tes as they are in water (13 CFU ml-1 after 24 h enrichment), but they do have the advantage of yielding information on V. vulnificus type as well as presence. Th e real-time PCR for the 4hr enrichment, which should have contained enough cells for detection, may have returned a false negative result due to the speed at which the samp les were centrifuged (16,000 X g). Analysis of the fractionation of the PCR pr oduct between the supernatant and pellet from replicate experiments showed that between 31% and 59% of the total DNA amplified appeared in the pellet (data not shown). A:B ratios and geographic distribution The A:B ratio observed for clinical isolates analyzed in this study was almost even at 19:17. These results contrast with those of previous studies, in which only 24% (8 of 34) of clinical isolates from oysters originating in the USA were type A or type AB (19, 30). However, six of the eight clinical cases caused by type A or AB V. vulnificus in the previous studies were contracted through the consumption of oysters harvested in Florida. In our study the proportion of type A clinical isolates from Florida wa s 11:12 (48% type A). The combined results of these studies [(current and 19, 30)] indicate th at Florida clinical isolates are much more likely to be type A than clinical isolates from other states, suggesting the possibility of geographical variation in the popul ation structure of V. vulnificus. A recent study explored relationships among V. vulnificus strains based on four distinct genetic measures (rep-PCR, 16S A/B, capsular polysaccharide operon group and RAPD profile) (7). Type A and AB clinical V. vulnificus strains were grouped in two distinct clusters (II and III) by rep-PCR pattern s with other oyster isolates, while the type

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50 B clinical isolates tended to be grouped separately in clusters I, IV and VII, supporting the hypothesis that genetically distinct subpopulations of V. vulnificus exist. In contrast to results for clinical isolates, the A:B ratios observed for environmental isolates from this study were in agreement with previous studies (19, 30); type A strains were the majority of those is olated from oysters collected from permitted harvest areas. Note, however, th at the frequency of type B V. vulnificus in oysters from prohibited areas was much greater than the fr equency of type B strains in oysters from permitted waters. The differences in strain distribution should be interpreted in light of the different sampling strategies employed in the studies; most of the environmental V. vulnificus strains typed in the previous studies (19, 30) were isolated from oysters collected from conditionally approved and commercially harvested sites, while the majority of the isolates for our study were isol ated from oysters in restricted areas with lower water quality. Furthermore, the majority of the environmental isolates for this study were collected between August and November while in previous studies comparing V. vulnificus type and isolate source (19, 30), isolat es were collected throughout the year. A study conducted in Galveston Bay, Texas noted seasonal variation in V. vulnificus A:B ratios, as type B isol ates were observed frequently in the warmer summer months, but not in cooler months (16). These data indicate th at factors such as s eason and water quality may influence A:B genotype ratios, and s hould be further explored. The typing of environmental and clinical V. vulnificus isolates from a broad geographic range, with a sampling strategy that accounts for seasonal va riation, and using a variety of molecular tools will be necessary to better understa nd the effects of envi ronmental and genetic variation on V. vulnificus infection rates.

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51 These rapid A/B PCR assays (~90 min in water samples) provide a quantitative piece of the puzzle about the population structure of V. vulnificus in coastal waters. These assays do not require culturing and c onfirming individual isolates, which is an added benefit over recentlypublished A:B typing methods (19, 30). This assay could prove useful in routine mon itoring of shellfish harvesting waters, which may in turn allow timely warning for consumers. Acknowledgements This project was funded in part by a grant from NOAA/UNH Cooperative Institute for Coastal and Estuarine Envi ronmental Technology, NOAA Grant Number(s) NA17OR1401 to VJH, and by Florida Sea Grant NA16RG-2195 R/LR-Q-26B to A.C. Wright and VJH. Cultures and technical a ssistance were kindly provided by Dr. D.V. Lim (USF), Dr. A. Cannons and C. Davis at USF Center for Biological Defense, R. Baker and R. Hammond at FDOH, J. Nordstro m at FDA, Dr. J.P. Gandhi at USF, Dr. A.C. Wright at the University of Florida and Dr. C. Tarr at the Centers for Disease Control. K. Burk and R. Gleeson of the Guana-Tolomato-Matanzas National Estuarine Research Reserve provided sampling advice and assistance. References 1. Acinas, S. G., L. A. Marcelino, V. Klepac-Ceraj, and M. F. Polz. 2004. Divergence and redundancy of 16S rRNA se quences in genomes with multiple rrn operons. J Bacteriol 186: 2629-2635. 2. Altschul, S. F., T. L. Madden, A. A. Schffer, J. Zhang, Z. Zhang, W. Miller, and D. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search pr ograms. Nucleic Acids Res 25: 3389-3402.

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52 3. Aznar, R., W. Ludwig, R. I. Amann, and K. H. Schleifer. 1994. Sequence determination of rRNA genes of pathogenic Vibrio species and whole-cell identification of Vibrio vulnificus with rRNA-targeted oligonucleotide probes. Int J Syst Bacteriol 44: 330-337. 4. Campbell, M. S., and A. C. Wright. 2003. Real-time PCR analysis of Vibrio vulnificus from oysters. Appl Environ Microbiol 69: 7137-44. 5. CDC. 1996. Vibrio vulnificus infections associated w ith eating raw oysters--Los Angeles, 1996. Morb Mortal Wkly Rep 45: 621-4. 6. CDC. 1993. Vibrio vulnificus infections associated with raw oyster consumption-Florida, 1981-1992. Morb Mortal Wkly Rep 42: 405-407. 7. Chatzidaki-Livanis, M., M. A. Hubbard, K. V. Gordon, V. J. Harwood, and A. C. Wright. 2006. Genetic distinctions among clinical and environmental strains of Vibrio vulnificus Appl Environ Microbiol. 72: 6136-6141. 8. Chen, C. Y., K. M. Wu, Y. C. Chang, C. H. Chang, H. C. Tsai, T. L. Liao, Y. M. Liu, H. J. Chen, A. B. Shen, J. C. Li, T. L. Su, C. P. Shao, C. T. Lee, L. I. Hor, and S. F. Tsai. 2003. Comparative genome analysis of Vibrio vulnificus a marine pathogen. Genome Res 13: 2577-2587. 9. DePaola, A., G. M. Capers, and D. Alexander. 1994. Densities of Vibrio vulnificus in the intestines of fish from the U.S. Gulf Coast. Appl Environ Microbiol 60: 984-8. 10. FAO Food and Nutrition paper. 2002. Risk assessment of Campylobacter spp. in broiler chickens and Vibrio spp. in seafood. Bangkok, Thailand. 11. Harwood, V. J., J. P. Gandhi, and A. C. Wright. 2004. Methods for isolation and confirmation of Vibrio vulnificus from oysters and environmental sources: a review. J Microbiol Methods 59: 301-16. 12. Jackson, J. K., R. L. Murphree, and M. L. Tamplin. 1997. Evidence that mortality from Vibrio vulnificus infection results from single strains among heterogeneous populations in shellfish. J Clin Microbiol 35: 2098-2101. 13. Kaper, J., H. Lockman, R. R. Colwell, and S. W. Joseph. 1979. Ecology, serology, and enterot oxin production of Vibrio cholerae in Chesapeake Bay. Appl Environ Microbiol 37: 91-103. 14. Kaysner, C. A., and A. DePaola. 2004. Vibrio cholerae V. parahaemolyticus V. vulnificus, and Other Vibrio spp., In Bacteriological Analytical Manual Online, 8th ed. Revision A, 1998. Chapter 9. Subs tantially rewritten and revised May 2004. http://www.cfsan.fda.gov/~ebam/bam-9.html

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53 15. Lee, S. E., S. Y. Kim, S. J. Kim, H. S. Kim, J. H. Shin, S. H. Choi, S. S. Chung, and J. H. Rhee. 1998. Direct identification of Vibrio vulnificus in clinical specimens by nested PCR. J Clin Microbiol 36: 2887-2892. 16. Lin, M., and J. R. Schwarz. 2003. Seasonal shifts in population structure of Vibrio vulnificus in an estuarine environment as revealed by partial 16S ribosomal DNA sequencing. FEMS Microbiol. Ecol. 45: 23-27. 17. Linkous, D. A., and J. D. Oliver. 1999. Pathogenesis of Vibrio vulnificus FEMS Microbiol Lett 174: 207-214. 18. Nicholas, K. B., H. B. J. Nicholas, and D. W. I. Deerfield. 1997. GeneDoc: analysis and visualization of genetic variation. EMBnet.news 4: 1-4. http://www.es.embnet.org/embnet_common/embnet.news/pdf/emnn42.pdf 19. Nilsson, W. B., R. N. Paranjype, A. DePaola, and M. S. Strom. 2003. Sequence polymorphism of the 16S rRNA gene of Vibrio vulnificus is a possible indicator of strain virulence. J Clin Microbiol 41: 442-446. 20. Panicker, G., and A. K. Bej. 2005. Real-time PCR detection of Vibrio vulnificus in oysters: comparison of oligonucleotid e primers and probes targeting vvhA. Appl Environ Microbiol 71: 5702-9. 21. Panicker, G., M. L. Myers, and A. K. Bej. 2004. Rapid detection of Vibrio vulnificus in shellfish and Gulf of Mexico water by real-t ime PCR. Appl Environ Microbiol 70: 498-507. 22. Perez-Rosas, N., and T. C. Hazen. 1989. In situ survival of Vibrio cholerae and Escherichia coli in a tropical rain forest wate rshed. Appl Environ Microbiol 55: 495-9. 23. Porter, K. G., and Y. S. Feig. 1980. The use of DAPI for identifying and counting aquatic microflora. Limnol. Oceanogr. 25: 943-948. 24. Rozen, S., and H. J. Skaletsky. 2000. Primer3 on the WWW for general users and for biologist programmers., p. 365-386. In K. S and M. S (ed.), Bioinformatics Methods a nd Protocols: Methods in Molecular Biology. Humana Press, Totowa, NJ. 25. Senoh, M., S. Miyoshi, K. Okamoto, B. Fouz, C. Amaro, and S. Shinoda. 2005. The cytotoxin-hemolysin genes of human and eel pathogenic Vibrio vulnificus strains: comparison of nucleotide sequences and application to the genetic grouping. Microbiol Immunol 49: 513-519. 26. Shapiro, R. L., S. Altekruse, L. Hu twagner, R. Bishop, R. Hammond, S. Wilson, B. Ray, S. Thompson, R. V. Tauxe, and P. M. Griffin. 1998. The role

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54 of Gulf Coast oysters harves ted in warmer months in Vibrio vulnificus infections in the United States, 1988-1996. Vibrio Working Group. J Infect Dis 178: 752-9. 27. Shiaris, M. P., A. C. Rex, G. W. Petti bone, K. Keay, P. McManus, M. A. Rex, J. Ebersole, and E. Gallagher. 1987. Distribution of indicator bacteria and Vibrio parahaemolyticus in sewage-polluted intertidal sediments. Appl Environ Microbiol 53: 1756-61. 28. Simpson, L. M., V. K. White, S. F. Zane, and J. D. Oliver. 1987. Correlation between virulence and colony morphology in Vibrio vulnificus Infect Immun 55: 269-272. 29. Takahashi, H., Y. Hara-Kudo, J. Miyasaka, S. Kumagai, and H. Konuma. 2005. Development of a quantitative real-tim e polymerase chain reaction targeted to the toxR for detection of Vibrio vulnificus J Microbiol Methods 61: 77-85. 30. Vickery, M. C., W. B. Nilsson, M. S. Strom, J. L. Nordstrom, and A. DePaola. 2007. A real-time PCR assay for the rapid determination of 16S rRNA genotype in Vibrio vulnificus J Microbiol Methods 68: 376-384. 31. Wang, S., and R. E. Levin. 2006. Rapid quantification of Vibrio vulnificus in clams ( Protochaca staminea) using real-time PCR. Food Microbiol 23: 757-61. 32. Wright, A. C., G. A. Miceli, W. L. Landry, J. B. Christy, W. D. Watkins, and J. G. Morris, Jr. 1993. Rapid identification of Vibrio vulnificus on nonselective media with an alkaline phosphatase-labeled oligonucle otide probe. Appl Environ Microbiol 59: 541-546. 33. Wright, A. C., and J. G. Morris, Jr. 1991. The extracellular cytolysin of Vibrio vulnificus : inactivation and relationship to virulence in mice. Infect Immun 59: 192-197. 34. Wright, A. C., L. M. Simpson, J. D. Oliver, and J. G. Morris, Jr. 1990. Phenotypic evaluation of acapsular transposon mutants of Vibrio vulnificus Infect Immun 58: 1769-1773.

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55 BOX-PCR GENOTYPING AND MULTILO CUS ANALYSIS OF VIRULENCE ASSOCIATED GENES OF ENVIRONMENTAL V. VULNIFICUS ISOLATES FROM PERMITTED AND PROHIBITED SHELLFISH HARVESTING AREAS Katrina V. Gordon and Valerie J. Harwood* Department of Biology, University of South Florida, Tampa, FL 33620 vharwood@cas.usf.edu

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56 Abstract Vibrio vulnificus is an opportunistic human pathogen that causes significant numbers of infections through recreational water contac t and consumption of raw seafood. Infections can range from gastroenteri tis to septicemia and death in 24 hours. In a recent study we noted a stat istically significant differen ce in the ratio of 16S rDNA types A, B and AB for V. vulnificus isolated from oysters harvested in permitted and prohibited shellfish harvesting areas, which are classified based upon the concentration of fecal indicator bacteria (fecal coliforms) in the water. Permitted shellfish harvesting areas yielded a higher proportion of t ype A isolates while prohibite d shellfish harvesting areas had a higher proportion of type B isolates. There was also a marginally higher number of type AB V. vulnificus isolated from the permitted area than the prohibited one. We hypothesized that this differen ce in proportion was due to environmental factor(s) that are correlated with shellfish water classificat ion. We also hypothesized that selective pressure exerted by environmental factors w ould be reflected in the genetic composition of V. vulnificus populations, which we assessed on a genomic level with BOX-PCR fingerprinting, and by assessing the relationshi p of various virulenc e-associated genes with the genomic patterns. Virulence-associat ed genes were assessed by PCR analysis of the vulnibactin ( viuB ) gene, 16S rRNA type and virulence correlated gene ( vcg ) to determine the genetic similarity among V. vulnificus isolated from permitted and prohibited shellfish harvesting areas. The 16S rDNA and vcg typing methods agreed as pr eviously noted, all type A isolates contained the vcgE gene sequence and all type B isolates contained the vcgC sequence. The viuB gene was present in only one type A/ vcgE isolate but in 42% of the

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57 type B/ vcgC isolates. Dendograms based on the BOX-PCR fingerprints of type A V. vulnificus separated strains into 3 clades, one of which was comprised mostly (80%) of isolates from permitted shellfish harvesting areas. These results demonstrate some genetic separation in the populat ions which is correlated w ith water quality in the oyster harvest area. Dendograms of th e BOX-PCR fingerprints of type AB isolates showed these were most similar to t ype A rather than type B isol ates. These results support the hypothesis that the population biology of V. vulnificus is influenced by the geographic region from which strains were isolated, and that the occurrence of certain genotypes may be affected by water quality. The high number of type B V. vulnificus containing the viuB gene isolated from prohibited shellfish ha rvesting areas adds credence to the use of fecal coliform concentrations to cl assify an area for oyster collection. Introduction Vibrio vulnificus is an opportunistic human pathogen that often causes infections through ingestion of undercooked seafood or c ontamination of open wounds. Symptoms of infection can be mild, such as gastro enteritis, but infection can quickly become systemic leading to septicemia and death in ~50% of cases (9, 29). Millions of individuals throughout the worl d eat raw oysters and come into contact with waters containing V. vulnificus, yet the rate of infection is ve ry low [less than 1 person in a million (4)]. Even with this low rate of infection, a recent Morbidity and Mortality Weekly Report (MMWR) surveillance summary of recreational water associated diseases and outbreaks showed V. vulnificus infections were responsible for the highest rate of hospitalization (82.7%) and death (12.8%) among the 47 Vibrio vulnificus infections

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58 reported (7). Using pulse field gel electrophoresis (PFGE), Jackson et al were able to determine that, although there were several strains present in an oyster, only one strain was responsible for infection (10). This data emphasizes the need to develop methods to assess the virulence potential of isolates and the relative risk associated with seafood consumption and recreational water usage. Several studies have attemp ted to define virulence fact ors of this pathogen which could be used to easily differentiate between virulent strains, thought to be most often involved in clinical infections, and less virulent strains, commonly found in the environment (5, 6, 14, 18, 22, 24, 26, 28). The correlation of V. vulnificus infection with diseases causing increased serum iron concentr ations have long been established (1). Wright et al. and Simpson and Oliver were also able to demonstrate the importance of iron availability through the ability of V. vulnificus to thrive in rabbit serum but human serum, which contains less freely available iron, was bactericidal (25, 35). The cytolytic activity of V. vulnificus has also been studied as a possi ble virulence factor. Chen et al identified the hlyIII hemolysin and showed that wit hout it, there was an increased LD50 in mice (6). In addition to identifying the actual genes involved in pathogenesis of V. vulnificus, work has been devoted to developing a ssays to differentiate strains based on virulence associated genes. Aznar et al identified the{Lee, 2004 #141} 17 nucleotide variations in the 16S rRNA gene of V. vulnificus strains separating them into two groups they termed type A and type B. These sequen ce variations would late r be used to develop a terminal restriction fragment length polymorphism assay (T-RFLP) (18), conventional PCR assays (13), and real-time PCR a ssays (8, 32) for differentiation of V. vulnificus

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59 isolates into these types. A second gene target, termed the virulence correlated gene ( vcg ) was identified by Warner et al (34). Rosche et al deve loped primers targeting this gene which classified 93% of the environmental and 72% of the clinical isolates tested as having the vcgE and vcgC sequence variations respectivel y (24). Genetic fingerprinting techniques have also been used to classify V. vulnificus strains based on virulence potential (10, 31). More recently, rep-PCR and multilocus sequence analysis targeting the CPS operon, 16S rRNA and vcg has been used to assess th e genetic similarity between clinical and environmental isolates. This me thodology was able to cla ssify isolates into clusters of highly similar strains having th e same virulence profiles (as assessed by the multilocus sequence analysis). Shellfish harvesting areas in Florida are monitored and classified by the Shellfish Environmental Assessment Section (SEAS) in the Bureau of Aquaculture Environmental Services ( http://www.floridaaquaculture.com/SEAS/SEAS_intro.htm ). Water samples at these areas are routinely analyzed to determ ine the fecal coliform (indicator organism) concentration, which is intended to reflect the risk of human pathogens of fecal origin in the water. Harvesting areas which are classifi ed as approved must meet the National Shellfish Sanitation Program (NSSP) 14/43 st andard (the fecal coliform median or geometric mean must not exceed 14 MPN/100 ml, and not more than 10 percent may exceed 43 MPN/100 ml). These areas are permitted to be commercially harvested. Harvesting areas classified as prohibited do not meet the NSSP 88/260 standard (the fecal coliform median or geometric mean must not exceed 88 MPN/100 ml, and not more than 10 percent may exceed 260 MPN/100 ml). Shellfish harvesting activities in these areas are prohibited due to actual or potentia l pollution and no oyste rs are obtained from

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60 these areas commercially. Ho wever, studies have found no correlation with fecal coliform concentrations and the presence of V. vulnificus (19, 20). In a recent study by Gordon et al (8) analyzing the 16S rRNA sequence of V. vulnificus isolates mostly from Florida oysters, a difference in the A:B ratio of isolates was seen based on shellfish harvesting area water quality. Isolates from retail oysters harvested from permitted, commercial shellfish harvesting areas in Apalachicola Bay, FL, had an A:B ratio of 10:1, in keeping with previous studies (18, 32). However the A:B ratio for isolates from areas that are not a pproved for shellfish harvesting was 5:8. There was no significant difference in the proportion of AB isolates found in each area. It is unclear whether seasonal variation in A/B or vcgE / vcgC type may have played a role in the difference in strain frequency seen. The Apalachicola isolates were collected in August when Gulf of Mexico water temperatur es are normally higher than October, when the majority of the Tampa isolates were obtained. Lin and Schwarz showed that decreasing water temperature favored an incr ease in the type B population (17) while Warner and Oliver recently showed that the vcgE population is always greater then the vcgC population in oyster tissue th roughout the year (33). Both these results contrast our previous findings for isolates from prohibited shellfish harv esting areas. The variability in observed ratios of type A/B frequencies in different studies led to an interest in the interand intra-population similarity between the V. vulnificus populations isolated from oysters harvested from permitted vs. prohibited shellfish harvesting areas. It was hypothesized that the environmenta l factors governing the classification of a harvesting area as permitted or prohibit ed, would influence the population biology and genotypes of the V. vulnificus inhabiting that area. In th is study, we utilized BOX-PCR

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61 fingerprinting of V. vulnificus harvested from oysters to determine the genetic similarity between strains (15). In add ition, the presence/absence of th e putative virulence factor vulnibactin ( viuB ) was determined and multilocus sequence analysis was carried out on the 16S rRNA and vcg gene. By combining the use of genetic fingerprinting with virulence associated genes it will be possible to get a more complete picture of whether parameters governing area classification pl aces a selective pressure on strains and therefore virulence potentials, of the isolates occupying that area. Methods Bacterial strains. Fifty five Vibrio vulnificus strains, obtained from our culture collection. These strains represented isolates from retail oysters (n= 23) harvested from permitted shellfish harvesting waters in Apalachicola, FL (Table 3) and isolates from oysters harvested from prohibited shellfish harvesting waters (n= 32) in Tampa Bay, FL and St. Augustine, FL (Table 4) in Florid a were used in this study. Fecal coliform concentrations were obtained from the Florida Department of Environmental Protection Storet Public Access Website ( http://storet.dep.state.fl.us /WrmSpa/default.do?page=home ). No fecal coliforms were detected at Apalachicola sites during August 2005 when the oysters were harvested; the highest concentration seen for that year was 23 MPN/100 ml. The mean fecal coliform concentrations at the Tampa Bay collect ion sites were 161.1 and 72.4 CFU/100 ml for July and October 2004 respectively, the two mont hs oysters were harvested. Oysters were collected, diluted 1:10 in alkaline peptone water (APW) and homogenized for 90 sec. The homogenates were then spread pl ated on selective media such as Vibrio vulnificus agar

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62 (VVA) or modified cellobiosepolymyxin B-colistin agar (mCPC) (12) with and without prior enrichment (incubation ove rnight at 37C). The plates were incubated overnight at 37C. Putative V. vulnificus isolates were confirmed us ing species specific primers targeting the vvhA gene (16). Six additional isolates were provided from the culture collection of the Food and Drug Administra tion (FDA) Gulf Coast Seafood Laboratory. These isolates were isolated from retail oysters harvested from permitted shellfish harvesting areas in Oregon, Florida, Texas and Louisiana. Isolates were grown overnight with shaking in brain heart infusion broth (BHI) supplemente d with 0.5% NaCl at 25C. DNA was extracted from these cultures us ing the DNeasy Tissue Kit (Qiagen) according to the manufacturers instruc tions, and quantified with the NanoDrop Spectrophotometer (Thermo Fisher Scientific MA USA). Strains were confirmed as V. vulnificus due to their amplification as type A or type B V. vulnificus primers. Strains MH1003-12 and ATCC 33814 were used as positive controls for type A/ vcgE / viuB -/ hlyIII and type B/ vcgC / viuB +/ hlyIII + PCR respectively.

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63 Table 3. Harvest area location and harvest date information along with results of multilocus analysis of isolates from permitted shellfish harvesting areas. Isolate Designation Harvest Location Harvest Date 16S rRNA type vcg type viuB F04 D15 Apalachicola, FL August 2005 A E F05 D06 Apalachicola, FL August 2005 A E F05 D08 Apalachicola, FL August 2005 A E F05 D14 Apalachicola, FL August 2005 A E F05 D16 Apalachicola, FL August 2005 A E F05 D23 Apalachicola, FL August 2005 A E F05 D24 Apalachicola, FL August 2005 A E F05 D25 Apalachicola, FL August 2005 A E F05 G02 Apalachicola, FL August 2005 A E F05 G10 Apalachicola, FL August 2005 A E F05 G27 Apalachicola, FL August 2005 A E F05 G29 Apalachicola, FL August 2005 A E F05 A06 Apalachicola, FL August 2005 A E F05 A20 Apalachicola, FL August 2005 A E F05 A30 Apalachicola, FL August 2005 A E F05 G06 Apalachicola, FL August 2005 A E Vv 99-609 Oregon February 1999 A E Vv 99-623 Florida December 1998 A E Vv 99-645 Texas May 1999 A E Vv 99-738 Florida April 1999 A E Vv 99-783 Louisiana May 1999 A E F05 G12 Apalachicola, FL August 2005 AB E F05 G18 Apalachicola, FL August 2005 AB E F05 G24 Apalachicola, FL August 2005 AB E F05 G26 Apalachicola, FL August 2005 AB E F05 D17 Apalachicola, FL August 2005 AB E F05 G05 Apalachicola, FL August 2005 B C F05 D04 Apalachicola, FL August 2005 B C + Vv 99-578 Louisiana November 1998 B C Multilocus PCR analysis. PCR analysis was used to assess the polymorphic sequences at primer anneali ng sites in the 16S rRNA and vcg genes as well as the presence/absence of the viuB gene. The DNA sequence at three polymorphic loci was determined by PCR analysis. PCR reactions were 25 l and contained 12.5 l GoTaq Green Master Mix (Promega, Madison WI), 0.5 M of appropriate prim er(s), and 2 l of the template

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Table 4. Harvest area location and harvest date information along with results of multilocus analysis of isolates from prohibited shellfish harvesting areas Isolate Designation Harvest Location Harvest Date 16S rRNA type vcg type viuB S1-2 Tampa, FL October 2004 A E S1-7 Tampa, FL October 2004 A E S1-8 Tampa, FL October 2004 A E S2-10 Tampa, FL October 2004 A E S2-11 Tampa, FL October 2004 A E S2-15 Tampa, FL October 2004 A E S3-5 Tampa, FL October 2004 A E S4-27 Tampa, FL October 2004 A E S4-48 Tampa, FL October 2004 A E + S7-3 Tampa, FL October 2004 A E MH1003-12 St Augustine, FL October 2003 A E MO0704-2 Tampa, FL July 2004 A E S6-13 Tampa, FL October 2004 A E S1-9 Tampa, FL October 2004 AB E S3-10 Tampa, FL October 2004 AB E S3-8 Tampa, FL October 2004 AB E S1-10 Tampa, FL October 2004 B C + S1-11 Tampa, FL October 2004 B C S1-14 Tampa, FL October 2004 B C S1-16 Tampa, FL October 2004 B C + S1-20 Tampa, FL October 2004 B C S2-20 Tampa, FL October 2004 B C + S2-22 Tampa, FL October 2004 B C S3-1 Tampa, FL October 2004 B C S3-16 Tampa, FL October 2004 B C + S3-4 Tampa, FL October 2004 B C S3-6 Tampa, FL October 2004 B C + S5-26 Tampa, FL October 2004 B C + S5-27 Tampa, FL October 2004 B C S5-28 Tampa, FL October 2004 B C + S6-14 Tampa, FL October 2004 B C S6-20 Tampa, FL October 2004 B C solution containing 10-50 ng DNA. The A/B type was determined for the 16S rRNA using primers previously developed and publis hed during the course of this dissertation research (8), VvAF1 (5-CAT GAT AGC TTC GGC TCA A-3) and VvAR1 (5-CAG CAC TCC TTC CAC CAT CAC-3) for t ype A and VvBF1 (5-GCC TAC GGG CCA 64

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65 AAG AGG-3) and VvBR1 (5-GTC GCC TCT GCG TCC AC-3) for type B. Separate PCR reactions were performed with each primer pair using the same thermal cycling profile; initial 5 min denaturation at 95 C, followed by 30 cycles of denaturation at 95C for 30s, annealing at 65C for 30s and exte nsion at 72C for 3 min followed by a final extension at 72 C for 7 min. The C/E type was determined for the vcg gene using primers P1 (5-AGCTGCCGATAGCGATCT-3) and P3 (5CGCTTAGGATGATCGGTG-3) for the vcgC gene sequence and P2 (5CTCAATTGACAATGATCT-3) and P3 for the vcgE type sequence (24). Thermal cycling profile was adjusted from publishe d temperatures to eliminate non-specific binding on the thermocyclers available. Initi al denaturation at 94C for 3 min followed by 30 cycles of amplification at 94C for 30s, 55C for 30s and 72C for 60s and final elongation of 72C for 2 min. The presence/absence of the vulnibactin gene ( viuB) was determined using primers F-viuB (5 -GGT TGG GCA CTA AAG GCA GAT ATA-3) and R-viuB (5-TCG CTT TCT CCG GGG CGG3) (21, 22). Thermal cycling profile was adjusted from published temperatures to eliminate non-specific binding on the thermocyclers available. Initial denaturation at 94C for 3 min followed by 30 cycles of amplification at 94C for 60s, 65C for 60s and 72C for 2 min and final elongation of 72C for 5 min. Five attempts were made to determine the presence/absence of the hlyIII gene using HF2 (5-CAT GTC GAC TAG CTG ACC ATT GCG-3) and HR2 (5-AAG GCA TGC GCT AAC TCA CCA GC-3) prim ers. The published thermal cycling protocol and four versions with modified annealing temperatures were used. Each traditional thermal cycling protocol contained an initial denaturation of 3 min at 95 C and 30 cycles of denaturation at 95C for 60s and extension at 72C for 60s and a final

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66 elongation at 72C for 2 min. Annealing temperat ures were as follows, a) 55C, b) 60C, c) 58C for 60s and d) 55C for 60s with th e addition of 2 l acetamide per PCR reaction to improve reaction specificity as this ther mal cycling profile. The fifth PCR attempt (e) was a touchdown format. There was an in itial denaturation at 94C for 3 min 30s followed by two cycles of 94C for 45s, and 65C for 45s, 72C for 30s, two cycles of 94C for 45s, and 64C for 45s, 72C for 30s, two cycles of 94C for 45s, and 63C for 45s, 72C for 30s, seven cycles of 94C for 45s, and 62C for 45s, 72C for 30s, and thirty cycles of 94C for 45s, and 55C for 45s, 72C for 30s, followed by a final elongation at 72C for 5 min. Amplified PCR reactions (5l) were load ed on a 2% agarose gel and run at 90V for 45 min in 1X TAE buffer with 1% ethi dium bromide solution (EtBr) (20 l/L). Digital images were captured with a FO TO/Analyst Investigator workstation (FOTODYNE, Hartland WI). PCR results were added into the data fields for each isolate in BioNumerics (Applied Maths Inc, Austin TX) to allow them to appear on the genotypic trees created with the BOX-PCR fingerprints (below). BOX-PCR genotyping. PCR reactions of 25 l were prepared using 12.5 l GoTaq Green Master Mix (Promega, Madiso n WI), 0.5 M of BO X A2R primer (15) and 2 l template DNA containing 10-50 ng DNA. Amplification was carried out using an initial 7 min denaturation at 95C followed by 35 cycles of denaturation at 90C for 30s, annealing at 40C for 60s and extension at 65C for 8 min., and a final elongation for 16 min at 65C. Fingerprint patterns we re visualized by agarose electrophoresis. Sixteen PCR reactions (5 l each) were run on one gel with four evenly spaced lanes containing 5 l 1 kb ladder (Promega, Madi son WI) and 3 l Blue/Orange 6X loading

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67 dye (Promega, Madison WI). Five microliters of each BOX-PCR reaction was placed on a 15 cm, 5mm thick 1.5% agarose gel (1.8g agarose, 120 ml 1X TBE buffer) run at 90V for 4 h. Gels were then stained at room temperature in 300 ml 1X TBE with 30 l of 1% EtBr for 20 min with shaking. The staini ng solution was then poured off and the gel was destained with diH2O at room temperature with shaking for 7 min. Digital images were entered into BioNumerics software (Appl ied Maths Inc, Austin TX). Bands in the ladder (3000 to 500 bp) were used as referen ce points to allow normalization across each gel and minimize the effect of inter/intra-ge l variation on the analys is of the patterns. After patterns were added to the software da tabase, relationships between isolates were assessed by constructing neighbor joining trees using the Pearson co rrelation for cluster analysis. Results and Discussion Multilocus typing. Thermal cycling protocols were able to produce the appropriately sized bands for the corres ponding gene targets for the 16S rRNA285bp for type A and 841bp for type B (Figure 1), vcg gene277 bp (Figure 2) and viuB gene504bp (Figure 3). The A/B genotype of the stra ins included below (Figure 1) is known because it was previously tested by A. DePaola at the FDA, and is included in the key (18, 32).

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Type B, 841 bp Type A, 285 bp 1500 1000 900 800 700 600 500 400 300 200 100 Key 1. 100 bp ladder A primers 2. Vv 99-578 (B) 3. Vv 99-738 (A) 4. Vv 99-783 (A) 5. Vv 99-645 (A) 6. PCR positive 7. PCR negative B Primers 8. Vv 99-578 (B) 9. Vv 99-738 (A) 10. Vv 99-783 (A) 11. Vv 99-645 (A) 12. PCR positive 13. PCR negative 1 2 3 4 5 6 7 8 9 10 11 12 13 Figure 1. Example of gel showing type A and type B V. vulnificus 16S rRNA PCR products. The genotype of each isolate is shown in parenthe ses in the key at right. A. B. 1 2 3 4 5 Key 1. 100 bp ladder 2. S1-14 3. S1-16 4. S1-20 5. S2-20 1 2 3 4 5 Key 1. 100 bp ladder 2. F05 G06 3. F05 G12 4. F05 G18 5. F05 G24 Figure 2. Example of gel showing A) vcgC and B) vcgE gene products. 68

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Key 1. 100 bp Ladder 2. F05 G06 3. F05 G12 4. F05 G18 5. F05 G24 6. F05 G26 7. F05 D17 8. F05 G05 9. F05 D04 10. Vv 99-578 11. S6-13 GD 12. S5-27GD 13. F05 G27 14. S4-48 15. Vv 99-783 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 Figure 3. Example of gel showing viuB gene products. 16. PCR positive 17. PCR negative Key L. 100 bp ladder 1. Vv ATCC 27562 2. MH1003-12 3. Vv 99-623 4. S1-9 5. S1-10 6. Vv 33814 7. Ent. faecium C68 All five attempts to produce only the 199bp hlyIII band were unsuccessful (Figure 4). A. 55C B. 60C C. 58C L 1 2 3 4 5 6 7 8 9 L 1 2 3 4 5 6 7 8 L 1 2 3 4 5 6 7 8 Figure 4. Examples of gels showing results for hlyIII PCR attempts. Panel A represents the attempt with annealing at 55C (results with conditions of 55C with acetamide and touchdown PCR were very similar); Panel B represents the attempt with an annealing temperature of 60C and Panel C represents the attempt at 58C. The arrows indicate the band of the correct size (~200 bp). 8. PCR negative 9. 50bp ladder 69

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70 The 16S rRNA and vcg sequence typing methods showed 100% agreement. Each isolate that contained the type A 16S rRNA gene also contained the vcgE gene sequence and those with the type B 16S rRNA gene also contained the vcgC sequence as previously demonstrated (5, 24), regardless of harvest area classification (Table 3 and 4). Those isolates that contained both type A and type B 16S rRNA sequences (type AB) also contained the vcgE gene sequence (but not the vcgC sequence). The co-occurrence of the AB type and vcgE has also been observed prev iously (5). Although one study reported that 84.6% of isolates containing both the vcgC and vcgE sequences were type AB isolates (33), no isolates observed to contain both the vcgC and vcgE sequences in this study. None of the 21 type A V. vulnificus isolates from permitted shellfish harvesting areas (permSHA) contained the viuB gene; however, one of the 13 type A isolates from prohibited shellfish harvesting ar eas (probSHA) did contain the viuB gene. No type AB isolates contained the viuB gene. One of the three type B isolates from permSHA did contain the viuB gene, as did 43.8% of the type B isolates from probSHA (Tables 3 and 4). Previous investigations into th e prevalence and distribution of the viuB gene have shown that 100% of vcgC type isolates contained the viuB gene as well (2, 22). The percentage of isolates showing a co-occurrence of vcgC and viuB here was lower than previously reported. BOX-PCR genotyping. The BOX-PCR protocol was able to produce reproducible patterns capable of differentia ting between strains. The B OX PCR fingerprints obtained for each isolate are detailed in Figures 5-9.

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Figure 5 Agarose gel #1 showing V. vulnificus BOXPCR patterns. The gel image has been cropped to show the portion of the pattern entered into BioNumerics and used to obtain similarity dendrograms. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 1617 18 19 20 Key 1. 1 kb ladder 2. F04 D15 3. F05 D06 4. F05 D08 5. F05 D14 6. F05 D16 7. F05 D23 8. 1 kb ladder 9. F05 D24 10. F05 D25 11. S1-2 12. S1-7 13. S1-8 14. 1 kb ladder 15. S2-10 16. S2-11 17. S2-15 18. S4-48 19. PCR negative 1 2 3 4 5 6 7 8 9 Key 1. 1 kb ladder 2. Vv 99-623 3. Vv ATCC 27562 4. S4-27 5. MH1003-12 6. S3-8 7. Vv 9031-96 8. PCR negative 9. 1 kb ladder Figure 6. Agarose gel #2 showing V. vulnificus BOX-PCR patterns. The gel image has been cropped to show the portion of the patt ern entered into BioNumerics and used to obtain similarity dendrograms. 71

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Figure 7. Agarose gel #3 showing V. vulnificus BOX-PCR patterns. The gel image has been cropped to show the portion of the pattern entered into BioNumerics and used to obtain similarity dendrograms. Figure 8. Agarose gel #4 showing V. vulnificus BOXPCR patterns. The gel image has been cropped to show the portion of the pattern entered into BioNumerics and used to obtain similarity dendrograms. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 Key 1. 1 kb ladder 2. S7-3 3. MO0704-2 4. S1-9 5. S3-10 6. S1-10 7. 1 kb ladder 8. S1-11 9. S1-14 10. S3-5 11. S6-13 12. F05 G02 13. F05 G27 14. 1 kb ladder 15. F05 G29 16. F05 A06 17. F05 A20 18. F05 A30 19. F05 G10 20. 1 kb ladder 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 Key 1. 1kb ladder 2. S1-16 3. S1-20 4. S2-20 5. S2-22 6. S3-1 7. S3-16 8. 1 kb ladder 9. S3-4 10. S3-6 11. S6-14 12. Vv 98-609 13. S6-20 14. 1kb ladder 15. S5-26 16. S5-27 17. S5-28 18. Vv ATCC 27562 19. Empty 20. 1kb ladder 72

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73 Figure 9. Agarose gel #5 showing V. vulnificus BOXPCR patterns. The gel image has been cropped to show the portion of the pattern entered into BioNumerics and used to obtain similarity dendrograms. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 Key 1. 1 kb ladder 2. Vv 99-645 3. Vv 99-738 4. Vv 99-783 5. F05 G06 6. F05 G12 7. 1 kb ladder 8. F05 G18 9. F05 G24 10. F05 G26 11. F05 D17 12. F04 G05 13. 1 kb ladder 14. F05 D04 15. Vv 99-578 16. S6-13 17. S5-27 18. Vv ATCC 27562 19. Empty 20. 1 kb ladde r The fingerprints were used to construct genotyp ic trees of each type (Figures 10 and 12) as well as combinations of two (Figures 11 and 13) or all types (Figure 14).

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100 80 60 Isolate F05 D16 F05 D24 F04 D15 F05 D14 F05 D06 99-623 S1-2 F05 D25 F05 G10 F05 D08 F05 D23 MH1003-12 S4-27 F05 G06 F05 A30 F05 A06 F05 A20 S1-7 S2-11 S2-15 F05 G02 Vv 98-609 S4-48 F05 G27 Vv 99-645 S1-8 MO0704-2 Vv 99-783 S7-3 F05 G29 Vv 99-738 S3-5 S6-13 S2-10 16S type A A A A A A A A A A A A A A A A A A A A A A A A A A A A A A A A A A vcg gene E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E viuB + I II III Figure 10. Dendrogram showing the similarity of type A V. vulnificus from permitted and prohibited shellfish harvesting areas. Three distinct clades were formed by the type A V. vulnificus isolates (Figure 10). Only 20% of the 15 isolates in clade I are from probSHA (S and M series isolates) while clades II and III are comprised of 45.5% and 62.5%, respectively, of V. vulnificus isolated from probSHA. The low percentage of isolates from probSHA in clade I suggests there is 74

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75 some genetic separation between isolates from these two types of harvesting areas. Understandably there is some similarity among isolates of different harvesting areas as well. This is represented by clades II and III, in which isolates from both harvesting areas are well-represented. The dendrogram of relationships amo ng both type A and type AB isolates separated the strains into three clades (Figure 11). Clade II contains the lowest frequency of isolates harvested from prohibited shellf ishing waters (3 out of 18 or 16%, again showing a distinction between populations isolat ed from permitted vs. prohibited waters. Clade II also contains the highest percentage of type AB isol ates (28% of total in that clade) compared to clades I and III, whic h are only 7% and 20% type AB isolates respectively.

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100 80 60 Isolate S1-2 S2-15 F05 G27 S2-11 S1-9 S1-7 F05 A06 F05 A20 F05 G02 Vv 98-609 S4-48 Vv 99-645 S1-8 F05 A30 F05 D25 F05 G10 F05 D08 F05 D23 MH1003-12 S4-27 S3-8 F05 G06 F05 G12 F05 G18 F05 D17 F05 D16 F05 D24 F04 D15 F05 D14 F05 D06 99-623 F05 G24 S7-3 F05 G29 Vv 99-783 MO0704-2 16S type A A A A A B A A A A A A A A A A A A A A A A B A A B A B A B A A A A A A A B A A A A vcg gen e E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E viuB + I S3-10 F05 G26 Vv 99-738 S3-5 S6-13 S2-10 A B A B A A A A E E E E E E II III Figure 11. Dendrogram showing the similar ity of type A and type AB V. vulnificus from permitted and prohibited shellfish harvesting areas. 76

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77 The dendrogram of type B V. vulnificus did not separate isolates into well-defined clades (Figure 12). Because there were only three type B isolates available from permSHA for testing, it is not pos sible to determine whether th ere is any separation of the BOX-PCR genotypes by harvest ar ea, although it has been established that the type B V. vulnificus strains are relatively rare in the oys ters harvested from permSHA. The few isolates from perSHA are not grouped together and they are only ~ 46% similar. Testing more type B isolates from perSHA would allow a better analysis of whether there is any separation of BOX-PCR genomic fi ngerprints by harvest area. As previously mentioned, a high percentage of type B/ vcgC isolates contained the viuB gene, which is consistent with other st udies (2, 22). However, possession of the viuB gene was not predictive of the relationship between strains based on BOX-PCR fingerprints; although two isolat es isolated from the same site (S3-6 and S5-26) that contained the viuB gene were ~96% similar (Figure 12).

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100 80 60 Isolate S5-28 F05 G05 S6-20 Vv 99-578 S1-16 S2-20 S3-1 S1-10 S2-22 S3-4 S6-14 S5-27 S3-6 S5-26 S1-20 F05 D04 S1-11 S1-14 S3-16 16S type B B B B B B B B B B B B B B B B B B B vcg gene C C C C C C C C C C C C C C C C C C C viuB + + + + + + + + Figure 12. Dendrogram showing the similarity of type B V. vulnificus from permitted and prohibited shellfish harvesting areas. When the genomic relationship of type B and type AB isolates was explored by BOX-PCR fingerprints (Figure 13), type AB isolates tended to cl uster together in clade I. They are ~57% similar to one another and to three of the type B is olates. This clade is also only ~46% similar to the other two clad es containing the majority of the type B isolates (n=16). 78

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100 80 60 Isolate F05 G18 F05 D17 F05 G12 S1-9 F05 G24 F05 G05 S1-16 S3-8 F05 G26 S3-10 S5-28 S2-20 S3-1 S1-10 S2-22 S3-4 S6-14 S5-27 S3-6 S5-26 S1-20 F05 D04 S1-11 S1-14 S6-20 Vv 99-578 S3-16 16S type A B A B A B A B A B B B A B A B A B B B B B B B B B B B B B B B B B B vcg gene E E E E E C C E E E C C C C C C C C C C C C C C C C C viuB + + + + + + + + I II III Figure 13. Dendrogram showing the similar ity of type B and type AB V. vulnificus from permitted and prohibited shellfish harvesting areas.. Type AB isolates cluster together when co mpared with type B isolates (Figure 13) but are interspersed in the de ndrogram with the type A isolates (Figure 10). This implies a higher level of similarity between type AB and type A V. vulnificus rather than type B V. vulnificus This could be an indication that type AB V. vulnificus have a lower virulence potential than type B isolates. 79

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100 80 60 Isolate S6-20 Vv 99-578 S7-3 F05 G29 Vv 99-783 MO0704-2 S3-10 F05 G26 Vv 99-738 S3-5 S6-13 S2-10 S1-2 S2-15 F05 G27 S2-11 S1-9 S1-7 F05 A06 F05 A20 F05 G02 Vv 98-609 S4-48 Vv 99-645 F05 D25 F05 G10 F05 D08 F05 D23 MH1003-12 S4-27 S3-8 F05 G06 F05 G12 F05 G18 F05 D17 F05 D16 16S type B B A A A A A B A B A A A A A A A A A B A A A A A A A A A A A A A A B A A B A B A B A vcg gene C C E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E E viuB + F05 D24 F04 D15 F05 D14 F05 D06 99-623 F05 G24 F05 A30 S5-28 S1-16 F05 G05 S1-8 S3-16 S2-20 S3-1 S1-10 S2-22 S3-4 S6-14 S5-27 S3-6 S5-26 S1-20 F05 D04 S1-11 S1-14 A A A A A A B A B B B A B B B B B B B B B B B B B B E E E E E E E C C C E C C C C C C C C C C C C C C + + + + + + + + Figure 14. Dendrogram showing the similarity of all V. vulnificus isolates. 80

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81 When all the isolates were included in one dendrogram, there was a clear separation between the types (Figure 14). Type A and type AB isolates grouped together except for two cases when type A isolates (FO5 A30 and S1-8) were most similar to a type B isolate (S5-28 and S3-16, respectively). The majority (84%) of the type B isolates were most similar to one another at an overa ll level of 60% simila rity. Two other type B isolates were most similar (68%) to one anot her than the closet branch containing type A and type AB isolates (56%). This trend of genetic distinctiveness between type A/ vcgE compared to type B/ vcgC isolates has been shown previous ly using clinical as well as environmental V. vulnificus strains isolated from permSHA throughout the U.S.; however, a different fingerprinting method (rep-PCR genomic typing) was used to establish the genetic relationships (5). It is promising that the BOX-PCR method determined similar relationships. The multilocus virulence gene analysis gave similar results to previous studies regardless of the fact that the majority of the isolates used in this study (90%) were obtained from harvest areas in Florida as opposed to throughout the U.S (2, 11, 24). Although other fingerprinting methods have been used for V. vulnificus and other gramnegative bacteria (3, 5, 10, 23, 31), this appear s to be the first publis hed account using the BOX-A2R primer to fingerprint V. vulnificus isolates. Other studies found no correlation between total V. vulnificus concentrations and fecal coliform concentrations (19, 20); however regulatory agencies continue to use this group of fecal indicator organisms as an indi cator of the health risk associated with consumption of seafood from shellfish ha rvesting areas. In th is study, the genetic separation seen in type A V. vulnificus associated with water quality, coupled with the

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82 presence of the vulnibactin gene in an isolate from a probSHA shows a possible correlation between water quality and virulence potential which should be investigated further. One study by Tamplin and Capers detailed growth of V. vulnificus in oysters and release of the bacterium into surrounding wate r (30). They found individual oysters could release between 105 to 106 V. vulnificus per hour. Therefore, another possibility for the variation in strain type seen at these two harvesting areas could be the large numbers of oysters present in Apalachicola oyster beds co mpared to Tampa. Differential growth rates of the two strains (27) could th en be magnified by the larger oyster beds releasing greater numbers of V. vulnificus thus leading to a larger incr ease in the type A population over the type B in Apalachicola and the revers e in Tampa. The high number of type B V. vulnificus containing the viuB gene isolated from prohibite d shellfish harvesting areas adds new credence to the classification methods used for shellfish harvesting areas. References 1. Blake, P. A., M. H. Merson, R. E. Weaver, D. G. Hollis, and P. C. Heublein. 1979. Disease caused by a marine Vibrio Clinical characteristics and epidemiology. N Engl J Med 300: 1-5. 2. Bogard, R. W., and J. D. Oliver. 2007. Role of iron in human serum resistance of the clinical and environmental Vibrio vulnificus genotypes. Appl Environ Microbiol 73: 7501-5. 3. Buchrieser, C., V. V. Gangar, R. L. Murphree, M. L. Tamplin, and C. W. Kaspar. 1995. Multiple Vibrio vulnificus strains in oysters as demonstrated by clamped homogeneous electric field gel el ectrophoresis. Appl Environ Microbiol 61: 1163-1168. 4. CDC. 1993. Vibrio vulnificus infections associated with raw oyster consumption-Florida, 1981-1992. Morb Mortal Wkly Rep 42: 405-407.

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83 5. Chatzidaki-Livanis, M., M. A. Hubbard, K. V. Gordon, V. J. Harwood, and A. C. Wright. 2006. Genetic distinctions among clinical and environmental strains of Vibrio vulnificus Appl Environ Microbiol. 72: 6136-6141. 6. Chen, Y. C., M. C. Chang, Y. C. Chuang, and C. L. Jeang. 2004. Characterization and virulence of hemolysin III from Vibrio vulnificus. Curr Microbiol 49: 175-9. 7. Dziuban, E. J., J. L. Liang, G. F. Craun, V. Hill, P. A. Yu, J. Painter, M. R. Moore, R. L. Calderon, S. L. Roy, and M. J. Beach. 2006. Surveillance for waterborne disease and outbreaks associ ated with recreational water--United States, 2003-2004. MMWR Surveill Summ 55: 1-30. 8. Gordon, K. V., M. C. Vickery, A. DePaola, C. Staley, and V. J. Harwood. 2008. Real-time PCR assays for quan tification and differentiation of Vibrio vulnificus strains in oysters and wate r. Appl Environ Microbiol 74: 1704-9. 9. Hlady, W. G., and K. C. Klontz. 1996. The epidemiology of Vibrio infections in Florida, 1981-1993. J Infect Dis 173: 1176-83. 10. Jackson, J. K., R. L. Murphree, and M. L. Tamplin. 1997. Evidence that mortality from Vibrio vulnificus infection results from single strains among heterogeneous populations in shellfish. J Clin Microbiol 35: 2098-2101. 11. Jones, M. K., E. Warner, and J. D. Oliver. 2008. Survival of and in situ gene expression by Vibrio vulnificus at varying salinities in estuarine environments. Appl Environ Microbiol 74: 182-7. 12. Kaysner, C. A., and A. DePaola. 2004. Vibrio cholerae V. parahaemolyticus V. vulnificus, and Other Vibrio spp., In Bacteriological Analytical Manual Online, 8th ed. Revision A, 1998. Chapter 9. Subs tantially rewritten and revised May 2004. http://www.cfsan.fda.gov/~ebam/bam-9.html 13. Kim, M. S., and H. D. Jeong. 2001. Development of 16S rRNA targeted PCR methods for the detectio n and differentiation of Vibrio vulnificus in marine environments. Aquaculture 193: 199-211. 14. Kim, Y. R., S. E. Lee, C. M. Kim, S. Y. Kim, E. K. Shin, D. H. Shin, S. S. Chung, H. E. Choy, A. Progulske-Fox, J. D. Hillman, M. Handfield, and J. H. Rhee. 2003. Characterization and pa thogenic significance of Vibrio vulnificus antigens preferentially expressed in septicemic patients. Infect Immun 71: 546171. 15. Koeuth, T., J. Versalovic, and J. R. Lupski. 1995. Differential subsequence conservation of inte rspersed repetitive Streptococcus pneumoniae BOX elements in diverse bacter ia. Genome Res 5: 408-18.

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84 16. Lee, S. E., S. Y. Kim, S. J. Kim, H. S. Kim, J. H. Shin, S. H. Choi, S. S. Chung, and J. H. Rhee. 1998. Direct identification of Vibrio vulnificus in clinical specimens by nested PCR. J Clin Microbiol 36: 2887-2892. 17. Lin, M., and J. R. Schwarz. 2003. Seasonal shifts in population structure of Vibrio vulnifcus in an estuarine environment as revealed by partial 16S ribosomal DNA sequencing. FEMS Microbiol. Ecol. 45: 23-27. 18. Nilsson, W. B., R. N. Paranjype, A. DePaola, and M. S. Strom. 2003. Sequence polymorphism of the 16S rRNA gene of Vibrio vulnificus is a possible indicator of strain virulence. J Clin Microbiol 41: 442-446. 19. Normanno, G., A. Parisi, N. Addante, N. C. Quaglia, A. Dambrosio, C. Montagna, and D. Chiocco. 2006. Vibrio parahaemolyticus, Vibrio vulnificus and microorganisms of fecal origin in mussels ( Mytilus galloprovincialis ) sold in the Puglia region (Italy ). Int J Food Microbiol 106: 219-22. 20. Oliver, J. D., R. A. Warner, and D. R. Cleland. 1983. Distribution of Vibrio vulnificus and other lactose-fermenting vibrios in the marine environment. Appl Environ Microbiol 45: 985-98. 21. Panicker, G., M. C. Vickery, and A. K. Bej. 2007. Erratum: Multiplex PCR detection of clinical and environmental strains of Vibrio vulnificus in shellfish. Can J Microbiol 53: 671. 22. Panicker, G., M. C. Vickery, and A. K. Bej. 2004. Multiplex PCR detection of clinical and envir onmental strains of Vibrio vulnificus in shellfish. Can J Microbiol 50: 911-22. 23. Parveen, S., K. M. Portier, K. Robinson, L. Edmiston, and M. L. Tamplin. 1999. Discriminant analysis of ribotype profiles of Escherichia coli for differentiating human and nonhuman sources of fecal pollution. Appl Environ Microbiol 65: 3142-7. 24. Rosche, T. M., Y. Yano, and J. D. Oliver. 2005. A rapid and simple PCR analysis indicates ther e are two subgroups of Vibrio vulnificus which correlate with clinical or environmenta l isolation. Microbiol Immunol 49: 381-389. 25. Simpson, L. M., and J. D. Oliver. 1983. Siderophore production by Vibrio vulnificus. Infect Immun 41: 644-9. 26. Simpson, L. M., V. K. White, S. F. Zane, and J. D. Oliver. 1987. Correlation between virulence and colony morphology in Vibrio vulnificus Infect Immun 55: 269-272. 27. Starks, A. M., K. L. Bourdage, P. C. Thiaville, and P. A. Gulig. 2006. Use of a marker plasmid to examine differential rate s of growth and death between clinical

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85 and environmental strains of Vibrio vulnificus in experimentally infected mice. Mol Microbiol 61: 310-23. 28. Stelma, G. N., Jr., A. L. Reyes, J. T. Peeler, C. H. Johnson, and P. L. Spaulding. 1992. Virulence characteristics of c linical and environmental isolates of Vibrio vulnificus Appl Environ Microbiol 58: 2776-82. 29. Tacket, C. O., F. Brenner, and P. A. Blake. 1984. Clinical features and an epidemiological study of Vibrio vulnificus infections. J Infect Dis 149: 558-61. 30. Tamplin, M. L., and G. M. Capers. 1992. Persistence of Vibrio vulnificus in tissues of Gulf Coast oysters, Crassostrea virginica exposed to seawater disinfected with UV light. Appl Environ Microbiol 58: 1506-10. 31. Tamplin, M. L., J. K. Jackson, C. Buchries er, R. L. Murphree, K. M. Portier, V. Gangar, L. G. Miller, and C. W. Kaspar. 1996. Pulsed-field gel electrophoresis and ribotype profiles of clinical and environmental Vibrio vulnificus isolates. Appl Environ Microbiol 62: 3572-3580. 32. Vickery, M. C., W. B. Nilsson, M. S. Strom, J. L. Nordstrom, and A. DePaola. 2007. A real-time PCR assay for the rapid determination of 16S rRNA genotype in Vibrio vulnificus J Microbiol Methods 68: 376-384. 33. Warner, E., and J. D. Oliver. 2008. Population structures of two genotypes of Vibrio vulnificus in oysters ( Crassostrea virginica) and seawater. Appl Environ Microbiol 74: 80-5. 34. Warner, J. M., and J. D. Oliver. 1998. Randomly amplified polymorphic DNA analysis of starved and viable but nonculturable Vibrio vulnificus cells. Appl Environ Microbiol 64: 3025-8. 35. Wright, A. C., L. M. Simpson, and J. D. Oliver. 1981. Role of iron in the pathogenesis of Vibrio vulnificus infections. Infect Immun 34: 503-7.

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86 RESEARCH SIGNIFICANCE The presence of V. vulnificus in shellfish and recreational waters is of grave health concern, particularly due to th e swift onset and severity of infections in predisposed individuals. Many studies have investigated the diversity of strains and virulence factors. The accumulated evidence indicates that the factors affecting virulence in V. vulnificus are very complicated and cannot be attributed to one specifi c gene product, as in other pathogenic bacteria (1, 9, 12, 20). Nevertheless, the poten tial virulence factors and virulence indicators identified (11, 14, 16, 24) have been used as targets to develop numerous assays to detect and to assess the virule nce potential of V. vulnificus strains from various environmenta l and clinical sources. Virulence Typing Methods The generally recognized method for detection of V. vulnificus in oysters and other seafood has been put forth by the FDA in the FDA Bacteriological Analytical Manual (FDA BAM) (8). The methodology advocated for the confirmation of isolates as V. vulnificus in the FDA BAM involves conventional PCR targeting the cytolysin/hemolysin, vvhA gene. This confirmation methodol ogy is cost effective and is relatively accessible to testing laboratories due to the lower cost of conventional versus real-time PCR therymocyclers. Other inve stigators have deve loped real-time PCR detection methods for the vvhA gene (3, 18); however, this gene is present in all V.

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87 vulnificus strains. It therefore simply confirms an isolates identity to the species level but offers no further indication of its virulence potential. Detection methods have been developed for virulence factors which have shown some involvement in virulence. These in clude the capsular polysaccharide (CPS) and vulnibactin gene ( viuB). A PCR assay has been develope d by Chatzidaki-Livanis et al. 2006 (5) which distinguishes between CPS alle les 1 and 2 which show some correlation to isolate origin and therefore virulence poten tial. Detection of the vulnibactin gene is now possible using a real-time PCR assay developed by Paniker et al. 2004(19). The sequence variations in the 16S rDNA have been targeted by several researchers allowing differentiation into type A and type B strains us ing different primers and/or probes. Kim and Jeong 2001 developed a tri-primer PCR assay to differentiate type A and type B strains (10) and Vickery et al developed a Ta qMan based real-time PCR assay (23). A PCR assay has also been developed for the virulence correlated gene ( vcg ) (21). All these methods allowing strain differ entiation have been tested with pure cultures and in some cases, th e ability to amplif y DNA from closely re lated Vibrios has been acknowledged by the investigators (10, 23 ). The real-time PCR assays developed in this dissertation work and published by Go rdon et al 2008 (6) have several major advantages over these previously published methods. The primer pairs have been shown to be specific for V. vulnificus and can therefore be used to detect V. vulnificus in water or oyster samples without the need to first obtain pure cultures. The water sample processing methodology actually bypasses any culturing whatsoever and allows large volumes of water to be rapidly tested. Du e to the use of primers and SYBR Green for

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88 real-time PCR detection the assay affords easy adaptation to conventional PCR if necessary. This work also provides a rapid, cost effective method for testing for the presence of V. vulnificus of varying virulence potentials to aid in the public health risk assessment of shellfish harvesting areas and recreational waters. Correlations with Virulence and Water Quality Past attempts to find correlations between V. vulnificus presence and fecal coliform concentrations were unsuccessful ( 15, 17). Both these stud ies determined the presence or absence of V. vulnificus through biochemical characteristics without taking into account virulence potential. However, f ecal coliform concentrations are still used in Florida to assess the quality of water bodies and therefore the risk associated with eating shellfish, especially raw shellfish, an d enjoying beaches and estuaries where V. vulnificus is normally found. During typing of numerous V. vulnificus strains it became apparent that there was a difference in strain proportions correlated with the sample site, i.e. whether isolates were obtained from a permitted (low fecal coliform concentrations) or prohibited (higher fecal coliform concentrations ) shellfish harvesting area. A higher percentage of type B strains was isolated from prohibited shellfis h harvesting areas than permitted shellfish harvesting areas. To investigate this phenom enon further, BOX-PCR fingerprinting was used to determine the genetic relatedness of these isolates. The seque nce variations in two loci correlated with virulence were also target ed as well as the presence or absence of the vulnibactin gene, to gain a bett er understanding of the virule nce potential of these isolates than one target would. These experiments showed that isolates from areas having higher

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89 fecal coliforms concentrations were more clos ely related to one anot her than those from areas having lower concentra tions that were permitted for shellfish harvesting. In addition, the vulnibactin gene which has been shown to directly impact virulence potential (13), was present in one of the type A/ vcgE (correlated with lower virulence potential) V. vulnificus isolated from an area of poor wa ter quality but none of the strains isolated from permitted shellfish harvesting areas. This shows that there is also the possibility for isolates having gene sequences correlated with lower virulence potential to be more virulent that those gene sequences would suggest. This could explain why there are some clinical isolates that are type A/ vcgE strains. This study is the first of my knowledge to use BOX-PCR fingerprinting to compare environmental isolates collected from areas of varying water quality. Howe ver, other methods have been used to determine the genetic fingerprints of V. vulnificus isolates in an attempt to determine the diversity in the environment (2, 7) as well as investig ate the relationship between environmental and clinical isolates (4, 22). Together these two studies have give n deeper insight into the ecology and population biology of V. vulnificus, both with respect to clinical vs. environmental source and possibly with respect to varying water quality. This work also provides a rapid, reproducible method which could feasibly be adopted by testing agencies in order to provide real-time information a bout the virulence potential of V. vulnificus in shellfish harvesting areas and recreational waters.

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90 References 1. Boyce, T. G., D. L. Swerdlow, and P. M. Griffin. 1995. Escherichia coli O157:H7 and the hemolytic-uremic syndrome. N Engl J Med 333: 364-8. 2. Buchrieser, C., V. V. Gangar, R. L. Murphree, M. L. Tamplin, and C. W. Kaspar. 1995. Multiple Vibrio vulnificus strains in oysters as demonstrated by clamped homogeneous electric field gel el ectrophoresis. Appl Environ Microbiol 61: 1163-1168. 3. Campbell, M. S., and A. C. Wright. 2003. Real-time PCR analysis of Vibrio vulnificus from oysters. Appl Environ Microbiol 69: 7137-44. 4. Chatzidaki-Livanis, M., M. A. Hubbard, K. V. Gordon, V. J. Harwood, and A. C. Wright. 2006. Genetic distinctions among clinical and environmental strains of Vibrio vulnificus Appl Environ Microbiol. 72: 6136-6141. 5. Chatzidaki-Livanis, M., M. K. Jones, and A. C. Wright. 2006. Genetic variation in the Vibrio vulnificus group 1 capsular polysaccharide operon. J Bacteriol 188: 1987-98. 6. Gordon, K. V., M. C. Vickery, A. DePaola, C. Staley, and V. J. Harwood. 2008. Real-time PCR assays for quan tification and differentiation of Vibrio vulnificus strains in oysters and wate r. Appl Environ Microbiol 74: 1704-9. 7. Jackson, J. K., R. L. Murphree, and M. L. Tamplin. 1997. Evidence that mortality from Vibrio vulnificus infection results from single strains among heterogeneous populations in shellfish. J Clin Microbiol 35: 2098-2101. 8. Kaysner, C. A., and A. DePaola. 2004. Vibrio cholerae V. parahaemolyticus V. vulnificus, and Other Vibrio spp., In Bacteriological Analytical Manual Online, 8th ed. Revision A, 1998. Chapter 9. Subs tantially rewritten and revised May 2004. http://www.cfsan.fda.gov/~ebam/bam-9.html 9. Keusch, G. T. 1998. The rediscovery of Shiga t oxin and its role in clinical disease. Jpn J Med Sci Biol 51 Suppl: S5-22. 10. Kim, M. S., and H. D. Jeong. 2001. Development of 16S rRNA targeted PCR methods for the detectio n and differentiation of Vibrio vulnificus in marine environments. Aquaculture 193: 199-211. 11. Lee, J. H., J. B. Rho, K. J. Park, C. B. Kim Y. S. Han, S. H. Choi, K. H. Lee, and S. J. Park. 2004. Role of flagellum and motility in pathogenesis of Vibrio vulnificus. Infect Immun 72: 4905-10.

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91 12. Levin, R. E. 2006. Vibrio parahaemolyticus a notably lethal human pathogen derived from seafood: A review of its pa thogenicity, characteristics, subspecies characterization, and molecular met hods of detection. Food Biotechnology 20. 13. Litwin, C. M., T. W. Rayback, and J. Skinner. 1996. Role of catechol siderophore synthesis in Vibrio vulnificus virulence. Infect Immun 64: 2834-8. 14. Nilsson, W. B., R. N. Paranjype, A. DePaola, and M. S. Strom. 2003. Sequence polymorphism of the 16S rRNA gene of Vibrio vulnificus is a possible indicator of strain virulence. J Clin Microbiol 41: 442-446. 15. Normanno, G., A. Parisi, N. Addante, N. C. Quaglia, A. Dambrosio, C. Montagna, and D. Chiocco. 2006. Vibrio parahaemolyticus, Vibrio vulnificus and microorganisms of fecal origin in mussels ( Mytilus galloprovincialis ) sold in the Puglia region (Italy ). Int J Food Microbiol 106: 219-22. 16. Okujo, N., M. Saito, S. Yamamoto, T. Yoshida, S. Miyoshi, and S. Shinoda. 1994. Structure of vulnibactin, a new pol yamine-containing siderophore from Vibrio vulnificus Biometals 7: 109-16. 17. Oliver, J. D., R. A. Warner, and D. R. Cleland. 1983. Distribution of Vibrio vulnificus and other lactose-fermenting vibrios in the marine environment. Appl Environ Microbiol 45: 985-98. 18. Panicker, G., M. L. Myers, and A. K. Bej. 2004. Rapid detection of Vibrio vulnificus in shellfish and Gulf of Mexico water by real-t ime PCR. Appl Environ Microbiol 70: 498-507. 19. Panicker, G., M. C. Vickery, and A. K. Bej. 2004. Multiplex PCR detection of clinical and envir onmental strains of Vibrio vulnificus in shellfish. Can J Microbiol 50: 911-22. 20. Reidl, J., and K. E. Klose. 2002. Vibrio cholerae and cholera: out of the water and into the host. FEMS Microbiol Rev 26: 125-39. 21. Rosche, T. M., Y. Yano, and J. D. Oliver. 2005. A rapid and simple PCR analysis indicates ther e are two subgroups of Vibrio vulnificus which correlate with clinical or environmenta l isolation. Microbiol Immunol 49: 381-389. 22. Tamplin, M. L., J. K. Jackson, C. Buchries er, R. L. Murphree, K. M. Portier, V. Gangar, L. G. Miller, and C. W. Kaspar. 1996. Pulsed-field gel electrophoresis and ribotype profiles of clinical and environmental Vibrio vulnificus isolates. Appl Environ Microbiol 62: 3572-3580. 23. Vickery, M. C., W. B. Nilsson, M. S. Strom, J. L. Nordstrom, and A. DePaola. 2007. A real-time PCR assay for the rapid determination of 16S rRNA genotype in Vibrio vulnificus J Microbiol Methods 68: 376-384.

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92 24. Warner, J. M., and J. D. Oliver. 1998. Randomly amplified polymorphic DNA analysis of starved and viable but nonculturable Vibrio vulnificus cells. Appl Environ Microbiol 64: 3025-8.

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ABOUT THE AUTHOR Katrina Gordon began her tertiary education at the University of the West Indies, Mona Campus in Jamaica where she earned a B. Sc. in Chemistry and Biochemistry. In 2002 she began as a Masters student in the De partment of Biology, at the University of South Florida (USF) and later switch ed to the Ph.D. program in 2004. At USF, she taught for the Cellular Bi ology, General Genetics and Determinative Bacteriology Laboratories. She was also a research assistant on grants involving the isolation and identification of Vibrio spp. as well as microbial source tracking. She presented her research at seve ral Southeastern branch (SEB) and General meetings of the American Society for Microbiology (ASM). She was awarded a travel grant to present at the 105th General meeting of the ASM, a student research grant from the SEB ASM and the Tharp Summer Research Fellowship fr om the University of South Florida, Department of Biology.