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Distribution and diversity of vibrio parahaemolyticus viruses (VpVs) and their hosts Comeau, Andreé Marcel 2005

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D I S T R I B U T I O N A N D D I V E R S I T Y O F Vibrio parahaemolyticus V I R U S E S ( V P V S ) A N D T H E I R H O S T S by A N D R E M A R C E L C O M E A U B.Sc.H., Acadia University, 1998 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE F A C U L T Y OF G R A D U A T E STUDIES (Microbiology and Immunology) THE UNIVERSITY OF BRITISH C O L U M B I A March 2005 © Andre Marcel Comeau, 2005 Abstract Vibrio parahaemolyticus viruses (VpVs) and their hosts were used as model organisms to explore the distribution and diversity of single-species populations in multiple habitats of coastal British Columbia. Abundances of host and virus were first monitored for a year in oysters. Temporal changes in host-range patterns, derived from bacterial and viral isolates, demonstrated that a significant shift in virus strains occurred during the winter months. Calculations of virus-induced mortality indicated that the presence of additional host species may be required to sustain viral production during summer. During winter, the persistent VpV population also required bacteria other than the host species and/or cells of altered culturability. Secondly, the distribution, phage-typing patterns, and genetic diversity of V. parahaemolyticus and total Vibrio spp. were examined in the water column and sediments. The bacteria were widespread and, not surprisingly therefore, geographic distribution was a poor determinant of strain diversity. Environment of isolation however, had a much more profound effect on genetic diversity and on phage-typing patterns to an even greater extent. Sources of high abundance produced strains with high susceptibility to viral infection; and the opposite was true for the water column. Thirdly, random amplification of polymorphic DNA was modified using a degenerate primer to produce unique and reproducible banding patterns from viral genomes. The findings described a rapid, PCR-based tool (DP-RAPD) for strain-typing viral isolates that allowed inferences to be made on genetic relatedness within groups of related viruses. Finally, the DP-RAPD was used in an investigation that examined the spatial variation in host-range and genetic diversity of newly isolated and characterized VpVs from the water column. VpVs were a nearly ubiquitous component of the water-column virioplankton, yet at very low abundances. When compared to oyster VpVs, it was clear that the major determinant of diversity was not geography, but the source habitat. Viruses from some of the farthest separated locations were closely related, while the diversity within some locations was very high. Overall, the diversity of viruses infecting a single Vibrio species was high and the host-virus populations were under similar phenotypic and genotypic selective pressures, mediated by the source habitat. ii Table of Contents Abstract ii Table of Contents iii List of Tables v List of Figures vi List of Abbreviations vii Preface viii Acknowledgements x Dedication xi Chapter I : Introduction to Marine Viruses and Methods of Assessing Their Impact in the Environment 1 1.1 Viruses in Marine Ecosystems 2 1.2 Determining the Impact of Marine Viruses 5 Chapter II : A Persistent, Productive, and Seasonally Dynamic Vibriophage Population Within Pacific Oysters (Crassostrea gigas) 9 2.1 Summary 10 2.2 Introduction 11 2.3 Materials and Methods , 13 2.4 Results 16 2.5 Discussion 25 Chapter III: Impact of Selection on the Distribution and Diversity of Vibrio spp. in Coastal Environments 31 3.1 Summary 32 3.2 Introduction 33 3.3 Materials and Methods 34 3.4 Results and Discussion 39 Chapter IV : The Use of Degenerate-Primed Random Amplification of Polymorphic DNA (DP-RAPD) for Strain-Typing and Inferring the Genetic Similarity Among Closely Related Viruses 55 4.1 Summary 56 4.2 Introduction 57 4.3 Materials and Methods 59 iii 4.4 Results and Discussion 61 Chapter V : Distribution and Diversity of Vibriophages in a Coastal Environment 68 5.1 Summary 69 5.2 Introduction 70 5.3 Materials and Methods 72 5.4 Results and Discussion 76 Chapter VI: Conclusions 90 6.1 Host-Virus Dynamics within Oysters 91 6.2 Host Distribution and Diversity within the Water Column and Sediments 92 6.3 Development of a Method to Assess Viral Diversity 93 6.4 Virus Distribution and Diversity within the Water Column 94 6.5 Conclusions and Contributions 95 References 98 List of Tables Table 2.1. Origins of bacterial strains used to determine the host range of VpVs 15 Table 2.2. The range of biological and physical parameters obtained from this study and others 18 Table 2.3. Pearson correlation analysis results of the physical and biological parameters measured in this study (from Figs. 2.1 and 2.2) 20 Table 2.4. Host range of VpVs isolated during summer 2000 and winter 2000/1 against co-occurring Vibrio parahaemolyticus (from 3 environments) and related Vibrio spp 21 Table 3.1. Total Presumptive Vibrio spp. (TPV) samples from coastal BC and corresponding ranges of measured values 34 Table 3.2. Bacterial strains (isolated in this study and others) used to determine host susceptibility and genetic fingerprinting patterns 38 Table 4.1. Selected characteristics of the virus strains used in this study 58 Table 5.1. Detection of VpVs in samples from the Strait of Georgia, BC 72 Table 5.2. Source of bacterial strains used to determine the host range of VpVs 75 Table 5.3. Characteristics of the VpV strains isolated in this study 79 Table 5.4. Host range of water-column VpV strains against co-occurring Vibrio parahaemolyticus from three environments and related Vibrio spp. (see Table 5.2) 83 Table 6.1. Chronological listing of all characterized Vibrio parahaemolyticus Viruses (VpVs) in the literature 96 v List of Figures Figure 1.1. Schematic of approaches used to analyze marine virioplankton 7 Figure 2.1. Physical data at the study site 17 Figure 2.2. Bacterial and viral abundances during May 2000 to May 2001 19 Figure 2.3. Total number of bacterial strains infected by each viral strain from oysters in summer (sensitivity) 22 Figure 2.4. Plating efficiency of summer versus winter oyster viral communities on strains of Vibrio parahaemolyticus from oysters in summer 24 Figure 2.5. Schematic representation of Vibrio parahaemolyticus-VpV dynamics in oysters. 28 Figure 3.1. Geographic distribution of samples 40 Figure 3.2. Malaspina system short-scale Vibrio spp. distribution 41 Figure 3.3. Distribution of Vibrio spp. in the Fraser River plume 43 Figure 3.4. Distribution of Vibrio spp. with depth 44 Figure 3.5. Distribution of Vibrio spp. within sediment cores 46 Figure 3.6. Vibrio spp. phenotypic diversity 48 Figure 3.7. Vibrio spp. ERIC-PCR fingerprinting 50 Figure 3.8. Vibrio spp. genotypic diversity 51 Figure 4.1. Analysis of algal virus genomes 63 Figure 4.2. Analysis of bacteriophage genomes 64 Figure 4.3. Influence of template mixtures on fingerprints 66 Figure 5.1. Geographic distribution of VpVs 77 Figure 5.2. Representative VpV morphologies 80 Figure 5.3. Phenotypic diversity of VpVs 84 Figure 5.4. VpV DP-RAPD fingerprinting -. 86 Figure 5.5. Genotypic diversity of VpVs 87 vi List of Abbreviations ATCC American Type Culture Collection BC British Columbia BCCDC British Columbia Centre for Disease Control CDC Centers for Disease Control and Prevention CFIA Canadian Food Inspection Agency CFU colony-forming units ChlV Chlorella Virus DNA deoxyribonucleic acid DNA pol DNA polymerase gene DP-RAPD degenerate-primed random amplification of polymorphic DNA dsDNA double-stranded deoxyribonucleic acid ERIC-PCR Enterobacterial repetitive-element intergenic consensus sequence PCR kb kilobases mer oligomer MLB Marine Luria-Bertani MPN most-probable number MpV Micromonas pusilla Virus PBS phosphate-buffered saline PCR polymerase chain-reaction PFGE pulse-field gel electrophoresis PFU plaque forming units psu practical salinity units SOG Strait of Georgia TBE Tris-borate EDTA TCBS thiosulfate citrate bile sucrose tdh thermostable direct hemolysin TPV total presumptive Vibrio spp. trh tdh-related hemolysin UPGMA unweighted pair group method with arithmetic mean VBNC viable but nonculturable VpV Vibrio parahaemolyticus Virus Preface This dissertation focuses on viruses of a single species in order to serve as a model for heterotrophic host-virus interactions in the marine environment. Virioplankton, numbering an estimated >10 viruses in the biosphere (Hendrix et al, 1999; Briissow and Hendrix, 2002), exert significant influence on marine phytoplankton and bacteria (reviewed in Wommack and Colwell, 2000). These effects include the reduction of productivity through viral-mediated mortality (Hewson et al, 2001; Wilhelm et al, 2002), the short-circuiting (virus shunt) of organic matter back into the dissolved pool through lysis products (Wilhelm and Suttle, 1999), mediating horizontal genetic exchange (Chibani-Chennoufi et al., 2004) and lysogenic conversion (Waldor and Mekalanos, 1996), and influencing the richness and evenness (i.e.: overall diversity) of host populations (Wommack et al, 1999b). It is the latter category of effects in which I have been most interested during the course of my dissertation - what can the distribution and the diversity of both populations tell us about the host-virus relationship? The layout of the thesis is as follows - an introductory chapter (I), followed by four research data chapters (ITV), and a final conclusions chapter (VI). So as to eliminate redundancy, the introductory chapter of the thesis will present only general information regarding viruses in the marine environment, methods of determining their impacts on ecosystems, and the objectives/aims of the thesis. The subsequent introductions within the four research chapters (II-V) will outline the specific background information pertinent to those sections. Chapters II-V are focused around investigations of host and virus in differing media (oyster, water column, and sediment), along with a novel method, in the following order - host and virus in oysters (II); host in water and sediments (III); a novel viral diversity method (IV); and virus in water and sediments (V). Chapter II is an exploration of the seasonal dynamics in abundance and phenotypic diversity of host and virus populations within oysters. Chapter III discusses the relationship between distribution and diversity, both phenotypic and genotypic, of water-column and sediment hosts from coastal BC. Chapter IV introduces a novel method for assessing the genetic diversity of viruses, which leads into the final research chapter (V) which examines the distribution and diversity, again phenotypic and genotypic, of viruses within the Strait of Georgia, BC. The final conclusions chapter (VI) summarizes the major findings of the thesis and places the significance of those findings within the context of marine viral ecology. Although all are to be submitted eventually, currently only chapter IV has been published in a peer-reviewed journal. It was published as: Andre M. Comeau, Steven M. Short, and Curtis A. viii Suttle. 2004. The use of Degenerate-Primed Random Amplification of Polymorphic DNA (DP-RAPD) for strain-typing and inferring the genetic similarity among closely related viruses. Journal of Virological Methods 118: 95-100. I am the primary author of this manuscript, and will be on those that follow from this dissertation work. As such, I was the author that designed and executed all experiments, and wrote the manuscripts. Steven M. Short provided theoretical discussions regarding the DP-RAPD assay and primer design, and was therefore included as second author on the manuscript resulting from the work in chapter IV. A small portion of the data {Vibrio parahaemolyticus abundances, n = 13; water salinities, n = 12; oyster temperatures, n = 13) from chapter II was provided by Enrico Buenaventura as part of a collaboration with the Canadian Food Inspection Agency (CFIA - Burnaby). As such, his name will appear as second author on the resulting manuscript. Curtis A. Suttle was (will be) the only other author listed on the manuscripts. He was my graduate supervisor and the principal investigator of the lab in which the research was conducted. As my graduate supervisor, he provided guidance for all of this work and was the primary editor of the manuscripts. hereby certify that the preceding authorship statements are correct. Curtis A. Suttle IX Acknowledgements First and foremost, I wish to thank Curtis for providing me with the opportunity to explore science in his lab. He has provided scientific guidance, "on-the-field and off, and has given me the freedom to express myself through my work, laying the foundation for a successful future career in science. My parents also deserve major thanks for their emotional and financial support throughout these many years of post-secondary education. While not always understanding what I do, they are always proud and supportive of my choices in life. Secondly, I would like to acknowledge helpful discussions with Alex, Alice, Amy, Andrew, CindyF, Emma, Janice, Jerome, Jessie, Karen, Sean, Steve, and Vera and that have helped formulate, and temper, my scientific reasoning throughout my doctoral experience. To my undergraduate students - Eric, Kelly, Daniel, and Luciana - thank you for allowing me to guide you along your path and, in the process, learn a little more about my journey. To all the other Suttle Lab members - I thank you for making the Lab a thoroughly enjoyable and never-boring environment within which to launch a scientific career! I would like to thank the members and crew, past and present, of the Suttle Laboratory, the CFJA (Burnaby), and the CCGS Vector for their help in sample collection throughout the years. I would like to gratefully acknowledge M. Kelly (BCCDC - Vancouver, BC), S. Short ("ex-Suttlelite"), H. Ackermann (Felix d'Herelle Reference Centre - Universite Laval, QC), E. Kutter (TESC - Olympia, WA), and J. Van Etten (UofN - Lincoln, NE) for providing strains and DNA; and to E. Buenaventura (CFIA - Burnaby, BC) for additionally providing Vibrio parahaemolyticus abundance data and biochemical identification of bacterial strains. I also deeply thank the W. Mohn Laboratory (and lab members; UBC) for the use of their gel analysis software; and A.M. Chan, J.E. Lawrence, and the UBC Biolmaging Facility for their assistance with electron microscopy. Lastly, this work was supported by a postgraduate fellowship from the Natural Sciences and Engineering Research Council of Canada (NSERC) and a travel grant from the UBC Faculty of Graduate Studies. x Dedication To my family, my ancestors, and my country - providers of my opportunities. Vive I 'Acadiel e:i Chapter I: Introduction to Marine Viruses and Methods of Assessing Their Impact in the Environment 1 1.1 Viruses in Marine Ecosystems In the late 1980s and early 1990s, it became apparent that high abundances of viruses existed in marine ecosystems (Bergh et al, 1989; Proctor and Fuhrman, 1990; Suttle et al, 1990). Coastal abundances are typically ca. 107 mL"1, with oligotrophic open-ocean abundances at ca. 106 mL"1 (reviewed in B0rsheim, 1993; Fuhrman and Suttle, 1993; Fuhrman, 1999; Wilhelm and Suttle, 1999; Wommack and Colwell, 2000). Viral abundances tend to be about 5- to 10-fold higher than that of prokaryotes and decrease with distance offshore and depth (Boehme et al, 1993; Cochlan et al, 1993; Jiang and Paul, 1994; Hara et al, 1996). It is now estimated that there are >1030 viruses in the biosphere (Hendrix et al, 1999; Briissow and Hendrix, 2002), representing the most abundant biologically active agents on Earth. As well as being abundant, viruses infecting many prokaryotic and eukaryotic species can be readily isolated from seawater (e.g. Moebus and Natkemper, 1983; Suttle et al, 1991; Kellogg et al, 1995). For example, viruses infecting the picoflagellate Micromonas pusilla and the cyanobacteria Synechococcus spp. have been found along the east and west coasts of North America, the coasts of Europe, in the Gulf of Mexico, and in the Mediterranean Sea (Cottrell and Suttle, 1991; Suttle and Chan, 1993; Waterbury and Valois, 1993; Wilson et al, 1993; Suttle and Chan, 1994; Cottrell and Suttle, 1995; Sahlsten, 1998; Zingone et al, 1999; Ortmann et al, 2002). Titers of individual, species-specific viruses have only been determined for a few hosts. For M. pusilla and the cyanobacterium Synechococcus, titers of viruses >105 mL"1 have been recorded (Suttle and Chan, 1994; Cottrell and Suttle, 1995). In contrast, maximum recorded abundances of viruses infecting Heterosigma akashiwo, a eukaryotic microalga, are ca. 104 L"1 (Chan et al, 1999) and for phage 3 1 infecting Vibrio parahaemolyticus, a marine heterotroph, are ca. 10 L" (Kellogg et al, 1995) in coastal waters. There have been few studies of marine sediments (Drake et al, 1998; Danovaro and Serresi, 2000), but all indications point to very high abundances (108-109 cc"1) with the presence of recoverable infectious viruses that may have been preserved for some time (Suttle, 2000; Lawrence et al., 2002). Temperate as well as lytic phages are frequently isolated which infect marine microbes (reviewed in Wommack and Colwell, 2000). However, induction of lysogens in marine environments appears to be responsible for only a small proportion of the total viral production in natural virus communities (Weinbauer and Suttle, 1996; Weinbauer and Suttle 1999; Williamson and Paul, 2004). At times, a significant portion of the prokaryotic community may be lysogenized (Jiang and Paul, 1996; Ortmann et al, 2002), but the primary significance of 2 lysogens may be to act as reservoirs of infective viruses and/or to mediate genetic exchange (Canchaya et al, 2003; Chibani-Chennoufi et al, 2004). There is strong evidence for high viral diversity in the marine environment. Although broad-host-range viral isolates exist (Matsuzaki et al, 1992; DePaola et al, 1998), most are species- or strain-specific (B0rsheim, 1993). Since titers of viruses infecting a host strain are frequently low, but the abundance of total viral particles is high, it implies the diversity of viruses is also high (Fuhrman and Suttle, 1993). Viruses have been isolated that infect most of the major classes of autotrophic and heterotrophic marine organisms. For the eukaryotes, viruses infecting coccolithophorids (Castberg et al, 2002), dinoflagellates (Tomaru et al, 2004a), diatoms (Nagasaki et al, 2004), other microalgae (Van Etten and Meints, 1999; Tai et al, 2003; Brussaard et al, 2004), and a heterotrophic nanoflagellate (Garza and Suttle, 1995) have been identified. For the prokaryotes, multiple phage have been isolated that infect Synechococcus spp. (e.g. Suttle and Chan, 1993), Prochlorococcus spp. (Sullivan et al, 2003), and many heterotrophs (e.g. Vibrio spp.; Kellogg et al, 1995). These results suggest that all marine organisms may be susceptible to viral infection (reviewed in Fuhrman and Suttle, 1993; Zingone, 1995). Although morphological diversity in natural viral assemblages is high, marine phage isolates tend to be 50-300 nm in length, contain dsDNA, and belong to one of three families of tailed phages (Myoviridae, Siphoviridae, or Podoviridae) (Ackermann et al, 1984; Frank and Moebus, 1987; Hara etal, 1991; Demuth etal, 1993). The genetic diversity of marine viral isolates is much greater than their morphological diversity. For example, various marker genes have been used to examine the genetic diversity of subsets of viruses in nature such as the Phycodnaviridae (Chen et al, 1996; Short and Suttle, 2002) and cyanomyoviruses (Zhong et al, 2002; Frederickson et al, 2003). Isolates of M. pusilla viruses, cyanophage, and vibriophage have also been examined using other molecular techniques (Cottrell and Suttle, 1991; Wilson et al, 1993; Kellogg et al, 1995). In all cases, the studies showed a high degree of genetic diversity and a widespread distribution of these virus-types in the world's oceans. Relatively recent genomic analyses of sequenced isolates and whole viral communities have reinforced these ideas. Seven marine tailed-phages have now been completely sequenced - roseophage SIOl (Rohwer et al, 2000), cyanophage P60 (Chen and Lu, 2002), and the vibriophages VHML (Oakey et al, 2002), VpV 262 (Hardies et al, 2003), KVP40 (Miller et al, 2003), and VP16C/VP16T (Seguritan et al, 2003) - all of which show distant relationships to known phages and significant amounts of novel sequence, indicating that the potential diversity of marine phage types is vast. Further confirming this are the first two 3 metagenomic studies conducted in the marine environment utilizing a shotgun-sequencing approach to characterize entire viral communities without the need for culturing. The analyses of pelagic (Breitbart et al, 2002) and marine sediment (Breitbart et al, 2004a) samples indicated that 65-75% of the sequence diversity was unknown and that the communities potentially contained >5000 viral types each. Studies on the abundance and diversity of marine viruses have generated interest in four major roles viruses play in the oceans (Fuhrman, 1999). The first role is to contribute to host cell mortality, including possible effects on primary production (reviewed in Suttle, 1994; Fuhrman, 1999; Wilhelm and Suttle, 1999). Initial results demonstrated that the simple addition of concentrated natural virus communities to natural phytoplankton communities led to significant reductions in primary productivity (Suttle et al, 1990; Suttle, 1992). Subsequent studies concluded that approximately 4-10% of the daily mortality of Synechococcus was the result of viral lysis (Suttle and Chan, 1994). The impact of viruses on heterotrophic bacterioplankton appears to be even greater. Although the value is highly variable, estimates indicate 10-20% of the bacterioplankton are lysed on a daily basis (Proctor and Fuhrman, 1990; Heldal and Bratbak 1991; Steward et al, 1992; Suttle, 1994; Fuhrman, 1999; Wilhelm et al, 2002). The second, yet related, role that viruses play is to affect biogeochemical cycling by short-circuiting the flow of carbon away from higher trophic levels and shunting nutrients into dissolved organic pools. Ecological models have now attempted to include the viral component in carbon, and other nutrient, cycles (Bratbak et al, 1990; Fuhrman, 1992; Wilhelm and Suttle, 1999). Recent estimates indicate that as much as 26% of the organic carbon produced in marine systems flows through the viral shunt (Wilhelm and Suttle, 1999). The last two roles of viruses in marine systems are to influence the diversity (richness/evenness) of individual populations and to mediate horizontal genetic exchange. These latter two present the greatest experimental challenges and, to-date, have not been explored in great detail (Weinbauer and Rassoulzadegan, 2004). Some attempts have been made with natural communities (Tarutani et al, 2000; Hewson et al, 2001), although most have focused on microcosms or batch cultures (Baross et al, 1974; Hennes et al, 1995; Jiang and Paul, 1998; Paul, 1999; Middelboe et al, 2001; Schwalbach et al, 2004; Winter et al, 2004b). These two areas of marine virus research should prove to be very fruitful in the future with the advent of novel technologies and the increasing use of whole-community genomic analysis. 4 1.2 Determining the Impact of Marine Viruses The past decade has produced significant knowledge regarding the impact of viruses on marine prokaryotic communities, but understanding of the effects on individual species is poor. There is little information on the dynamics of specific host cell populations or of the specific viral populations infecting them. There is also little known about the diversity of host and viral populations and the factors which influence or control them. The overall impact of viruses on the mortality and diversity of host populations is therefore unknown. There are various approaches to the study of marine virus communities, and specific virus populations, most of which either depend upon direct -methods, nucleic acid techniques, or culture-based approaches that bridge the former two (Fig. 1.1). 1.2.1 Direct Methods Direct methods of analyzing the virioplankton were the first to be employed and give information primarily on the mortality caused by viruses. Determinations of the abundances and distributions of hosts and virioplankton have benefited from the application of epifluorescent microscopy (Hennes and Suttle, 1995; Noble and Fuhrman, 1998) and flow cytometry (Marie et al, 1999). These data can be compared to physical and other biological parameters to determine which factors are correlated to measured abundances. Host and virus abundance data also allow the determination of contact (encounter) rates (Murray and Jackson, 1992), or the rate at which collisions occur between hosts and viruses. The contact rates indicate the maximum rate of infection possible. These values can be compared to other direct measurements of the virioplankton. Various studies have looked at rates of virus production (Wilhelm et al, 2002; Evans et al, 2003; Winter et al, 2004a), virus decay and repair (Heldal and Bratbak, 1991; Suttle and Chen, 1992; Noble and Furhman, 1997; Weinbauer et al, 1997; Wilhelm et al, 1998), reduction of production under the influence of concentrated viral communities (Suttle et al, 1990; Hewson et al, 2001), or have attempted to determine the proportion of visibly infected cells in prokaryotic communities (Proctor et al, 1993; Hennes and Simon, 1995) to obtain information, with the help of mathematical models (Binder, 1999; Middelboe, 2000; Coombs et al, 2003), on the fraction of mortality due to viral lysis. The role of lysogeriy in mortality and production (Jiang and Paul, 1994; Jiang and Paul, 1996; Weinbauer and Suttle, 1996; Weinbauer and Suttle, 1999) can be estimated using various inducing agents (mitomycin C, U V , heat); however its affect on lateral gene transfer, as mentioned previously, is probably of greater 5 importance (reviewed in Canchaya et al, 2003). Generally, these measurements of mortality are difficult to conduct for specific populations (i.e.: a single species), therefore other methods must be used to determine the effect of viruses on specific populations. 1.2.2 Nucleic Acid Methods Whereas the previously mentioned direct methods generally assess the mortality caused by viruses, the isolation and analysis of virioplankton DNA/RNA allows for the examination of genetic diversity. Methods of assessing whole-community diversity have generally been restricted to the use of pulse-field gel electrophoresis (PFGE) to examine genome size distributions and variability (Wommack et al, 1999a; Wommack et al, 1999b; Steward et al, 2000; Jiang et al, 2003) or the more recently applied metagenomic approach of shotgun sequencing entire virus assemblages from a few samples (Breitbart et al, 2002; Breitbart et al, 2004a). Moving to the more specific, most culture-independent diversity studies have employed a single-gene PCR approach, generally followed by separation using denaturing gradient gel electrophoresis (DGGE) and/or sequencing, to examine the diversity of a particular family or subset of viruses. Examples of these are the use of the DNA polymerase for the phytoplankton-infecting Phycodnaviridae (Chen et al, 1996; Short and Suttle, 2002) and Podoviridae (Breitbart et al, 2004b), the capsid gene (gp20) for cyanomyoviruses (Zhong et al, 2002; Frederickson et al, 2003), and the RNA-dependant RNA polymerase (RdRp) for picorna-like viruses (Culley et al, 2003). Although these methods allow for the examination of the whole community or can be targeted to specific types of viruses (e.g. cyanomyoviruses), they cannot be applied to a subset of viruses infecting one host species since there are no host-specific markers amongst the many different types of viruses. The application of an isolation step must therefore be employed to arrive at this level. 6 Direct Enumeration Virus Production Decay/Repair Lysogeny (LGT) Single-Type Distribution Host-Range/Typing Genomics Transduction (LGT) DNA/RNA Single-Gene PCR PFGE Hybridization Metagenomics (LGT) I Diversity - L = Figure 1.1. Schematic of approaches used to analyze marine virioplankton. Approaches are categorized as to whether they are direct (left) or nucleic acid (right) methods of analyzing whole-community samples, or culture-based (centre) methods targeting species-specific viruses. Direct methods generally lead to information on mortality, nucleic acid methods lead to information on diversity, and culture-based methods give both types of information. Methods that also give information on lateral gene transfer (LGT) are indicated. 7 1.2.3 Culture-Dependent Approaches Culture-dependent approaches serve as a link between the two former methods, providing access to information on both mortality and diversity. The isolation and/or enumeration of viruses infecting a single species is generally conducted by screening concentrated virus community samples (Suttle et al, 1991) by plaque assay (double-layer agar overlay) or by liquid enrichment cultures using the host of interest (Suttle, 1993). Once clonal isolates have been obtained, a myriad of characterization and experimental studies can be carried out to determine the impact of these viruses. Some of these include ultra-structural studies (Frank and Moebus, 1987), classification schemes (Ackermann et al., 1984), one-step growth experiments (Proctor et al, 1993), protein characterizations (Matsuzaki et al, 1998), UV/chemical sensitivities (Koga et al, 1982; Kellogg and Paul, 2002), microcosm (Hennes et al, 1995; Middelboe, 2000) and transduction (Baross et al, 1974; Muramatsu and Matsumoto, 1991; Jiang and Paul, 1998) experiments, and many genetic measures. Host-range and genetic diversity of the viral and host isolates can also be used to describe phage-host dynamics. The host ranges of viral isolates, and the converse patterns of host sensitivities (phage-typing), can be used as a measure of diversity, indicating for example whether hosts/viruses exhibiting different host ranges are separated geographically or temporally (Moebus and Nattkemper, 1981; Moebus and Nattkemper, 1983; Tomaru et al, 2004b), or whether particular subsets are more resistant or susceptible in different locations or times of the year (Waterbury and Valois, 1993; Suttle and Chan, 1994; Garza and Suttle, 1998). The genetic diversity of hosts can be measured in various ways, given the existence of rRNA or other markers, which are appropriate for the organism in question. Similarly, PCR analysis of viral markers can be used if there is a priori information regarding the virus type under examination. However, the assessment of viral diversity has generally been restricted to restriction digests of genomic DNA (Cottrell and Suttle, 1991; Wilson et al, 1993) and DNA/DNA hybridizations (Kellogg et al, 1995; Wichels et al, 1998; Matsuzaki et al, 2000) for completely novel viruses. The new technique of degenerate-primed random amplification of polymorphic DNA (DP-RAPD) will be introduced in this dissertation (Chapter IV) as an easier and more universal alternative. Finally, whole-genome sequencing can be used to gain insights into the diversity and evolution of individual marine viruses (Rohwer et al, 2000; Chen and Lu, 2002; Hardies et al, 2003). All of these culture-dependent techniques have inherent bias, but it is unavoidable in a selective system which is the only current way to assess the impact of viruses infecting a single species. 8 Chapter II: A Persistent, Productive, and Seasonally Dynamic Vibriophage Population Within Pacific Oysters (Crassostrea gigas) 2.1 Summary Most studies in marine viral ecology have focused on the pelagic environment, even though substantial populations of viruses and host cells are within sediments and shellfish. Abundances of Vibrio spp. and viruses infecting Vibrio parahaemolyticus (VpVs) were monitored for a year in Pacific oysters and water collected from Ladysmith Harbour, BC. Bacteria were highly seasonal, whereas high titers of VpVs (0.5-11 x 104 viruses cc"1) were persistent year-round in oysters, even when V. parahaemolyticus was undetectable (<3 cells cc"1). Viruses were not detected (<10 mL"1) in the water column. Host-range studies on 13 viral isolates demonstrated that VpV strains could infect most V. parahaemolyticus strains from oysters and sediments (62 and 74% of the total 91 and 65 host-virus combinations, respectively), but few water-column strains (30% of 91 combinations). Ten of the 13 viruses also infected more than one species among V. alginolyticus, V. natriegens, and V. vulnificus. As winter approached and potential hosts disappeared, the number of host strains that the viruses were capable of infecting decreased (-50% reduction) and, in the middle of winter, the VpV community could no longer be plated very efficiently (14% of total PFUs) on summer host strains. Estimations of virus-induced mortality on V. parahaemolyticus demonstrated that the presence of additional host species, or additional undetected/unculturable host cells, may be required to sustain viral production within oysters during summer. During winter, the persistent VpV population also required bacteria other than the putative host species (since they were undetectable) and/or they continued to replicate on cells that were no longer culturable. 10 2.2 Introduction Over the past decade, much work has examined the effects of viral infection on marine microbial communities (reviewed in Suttle, 1994; Fuhrman, 1999; Wilhelm and Suttle, 1999; Wommack and Colwell, 2000). Some examples are the impact of lytic viruses on primary production (Suttle et al, 1990; Suttle, 1992; Hewson et al, 2001) and bacterial mortality (Proctor and Fuhrman, 1990; Heldal and Bratbak, 1991; Steward et al, 1992; Wilhelm et al, 2002), and the effects of induction of temperate phage on microbial mortality (Jiang and Paul, 1994; Jiang and Paul, 1996; Weinbauer and Suttle, 1996; Weinbauer and Suttle, 1999; Ortmann et al, 2002). These investigations have largely focused on pelagic environments, even though the abundances of bacteria and phage can be much higher in other environments. For example, Vibrio spp. and vibriophages are common in the marine environment (B0rsheim, 1993; Thompson et al, 2004a), are easily culturable (Eilers et al, 2000), and have served as model host-virus systems in many water-column investigations (e.g. Suttle and Chen, 1992; Proctor et al, 1993; Hennes et al, 1995; Weinbauer et al, 1997; Long and Azam, 2001; Kellogg and Paul, 2002; Thompson et al, 2004b). However, Vibrio spp. abundances are typically much higher in sediments (104 g"1; Pfeffer et al, 2003), plankton (109 g"1; Kaneko and Colwell, 1978), and shellfish (105 g"1; Arias et al, 1999) than in the water column (-10 mL"1; Cavallo and Stabili, 8 9 -2004). In contrast, even though virus particles are extremely abundant in sediments (10 -10 cc" '; Drake et al, 1998; Danovaro and Serresi, 2000) and in the water column (~107-108 mL"1; 5 8 1 Wommack and Colwell, 2000), vibriophage are most abundant in molluscs (10 -10 g" ; Baross et al, 1978b; DePaola et al, 1998), relatively rare in the water column (~2 mL"1; Kellogg et al, 1995), and frequently undetectable in sediments. The gastrointestinal pathogen Vibrio parahaemolyticus, acquired through the consumption of raw shellfish, has been of interest in coastal BC since it was responsible for a major outbreak of disease in 1997 (Centers for Disease Control and Prevention, 1997). High abundances of V. parahaemolyticus (104 g"1; DePaola et al, 2000), and the viruses which infect it (VpVs; 106 g"1; Baross et al, 1978b) occur in oysters. Yet, with the exception of a study showing transduction of the agarase gene between V. alginolyticus and V. parahaemolyticus (Baross et al, 1974), and host-range studies using V. vulnificus (DePaola et al, 1997; DePaola et al, 1998), little work has been done in this environment. The high abundances in oysters of V. parahaemolyticus and the phage that infect it provided an excellent opportunity for examining phage-host interactions in a non-pelagic marine environment. 11 This study examined seasonal changes in the abundances of V. parahaemolyticus, its phage, total presumptive Vibrio in oysters and the neighboring water column, as well as host range and sensitivity of host and virus strains. This study shows that oysters are likely the primary source of viruses infecting V. parahaemolyticus in oysters and in the water column. Furthermore, seasonal shifts in patterns of host range provide strong evidence that the composition of the virus community changes during winter, and that the persistence of these viruses in oysters during winter is supported by hosts other than V. parahaemolyticus. These results have implications for other pelagic host-virus systems that also have high abundances in other environments and/or that respond to strong seasonal trends. Production of many other marine viruses may depend on inputs from non-water-column sources and contributions from hosts beyond the primary species. 12 2.3 Materials and Methods 2.3.1 Sample Collection Oyster and seawater (250 mL) samples were collected from a commercial aquaculture bed in Ladysmith Harbour, BC, Canada (49°00'N, 123°49'W) biweekly from June to September 2000 and approximately monthly thereafter until May 2001. Samples for plating efficiency tests (see below) were also collected in August 2003. Seawater samples were collected adjacent to the oysters, and all samples were transported on ice and analyzed within 24 h. Oyster samples were shucked and homogenized (US Food and Drug Administration, 1998) before analysis, and comprised 10-12 pooled individuals collected across the bed, except for August 2003 in which 10 oysters were analyzed separately and then pooled for further testing. 2.3.2 Physical Properties Temperature within the oyster was taken, at time of collection, using a hand-held thermometer and water salinity was determined using a hand-held refractometer (both Fisher Scientific). 2.3.3 Total Presumptive Vibrio spp. (TPV) Enumeration and Isolation Oyster homogenates were vortexed and centrifuged (4100 x g for 60 s) to sediment large debris and the supernatants sub-sampled. While centrifugation may slightly reduce bacterial abundance (-10% in test samples), filtration was not possible without prior clarification. Filtration was the desired method given its greater precision (e.g.: accepted standard for water quality) over methods such as spread plating. To obtain estimates of mesophilic TPV abundance, water and oyster sub-samples were serially diluted in duplicate 10 mL (final volume) of ultrafiltered (30 kDa cut-off, Millipore; Suttle et al., 1991), sterilized natural seawater diluted to 15 psu with reverse-osmosis water (Millipore). The dilutions were filtered onto 0.45-pm pore-size cellulose membranes (47 mm HAWP, Millipore) which were laid, two per plate, onto thiosulfate citrate bile sucrose (TCBS) agar (BD Gibco). The plates were incubated at 25°C for -24 h. Colonies were purified by serially re-streaking 3 times onto Marine Luria-Bertani agar plates (MLB; 0.5 g L"1 each of casamino acids, peptone, and yeast extract, 0.3% v/v glycerol, 1% w/v agar, in 15 psu ultrafiltrate base), and identified biochemically (API 20 E®, bioMerieux). 13 2.3.4 Vibrio parahaemolyticus Enumeration and Isolation Abundances of V. parahaemolyticus were determined in oysters as outline by the US Food and Drug Administration (1998). Oyster homogenates were diluted 10-fold in 2% NaCl and inoculated into a series of alkaline peptone water (APW) tubes for an overnight MPN assay. Positive tubes were plated onto TCBS agar and colonies of V. parahaemolyticus were confirmed biochemically (API 20 E®). 2.3.5 Vibrio parahaemolyticus Virus (VpV) Enumeration and Isolation The titers of VpVs were determined in oyster and water samples by plaque assay (Suttle, 1993) using MLB plates and top agar overlay (0.6% w/v agar). In order to confirm that the pooled oyster samples were representative of individuals, 10 oysters were titered for VpVs separately, then as a pooled sample. The pooled sample titer was slightly lower, but within one standard deviation of the mean obtained from the individuals. Duplicate parallel dilutions were performed using the same diluent as for TPV counts. The host strain used for enumeration, clinical isolate 94Z944 El-80 (British Columbia Centre for Disease Control), was determined to be the most permissive host in terms of total viral abundance in test assays and is herein referred to as the control strain. Although this may bias the types of viruses recovered, hopefully the widest range possible will be sampled. For most samples, 1-3 different viruses (indicated by plaque morphology) were plaque-purified for 3 rounds to provide clonal strains. Viruses were then amplified by the plate-lysate method (elution of phage from a lawn of confluent lysis; Sambrook et al., 1989), 0.22 pLm-filtered, and maintained at 4°C in the dark. 2.3.6 Host Range and Plating Efficiency Assays The host ranges of viral strains were determined using strains of V. parahaemolyticus and other Vibrio spp. that were obtained during this study or from other sources (Table 2.1). Ten to 20 pi of amplified virus stocks were spotted into lawns of bacteria and monitored for zones of clearing for 1-7 d. Two oyster homogenates, from 21 February 2001 (the "winter" community) and from 28 August 2003 (the "summer" community), were used to compare the relative plating efficiencies among several summer strains of V. parahaemolyticus. Plaque assays and efficiencies were expressed relative to the control strain. 14 2.3.7 Statistical Analysis Undetectable values were assumed to equal the detection limit for calculations of mean abundances and for correlation analyses. Correlations were performed using VassarStats (http://faculty.vassar.edu/lowry/VassarStats.html). Plaque-forming units (PFU cc"1) were considered representative of the abundances of viruses (viruses cc"1) infecting V. parahaemolyticus, while the number of colony-forming units (CFU mL"1 or cc"1) and most-probable-number (MPN cc"1 for V. parahaemolyticus counts) were taken to be indicative of the bacterial cell abundances (cells mL"1 or cc"1). Bacterial and viral abundances in the oysters were expressed per cubic centimetre of tissue homogenate. Table 2.1. Origins of bacterial strains used to determine the host range of V p V s . All strains were isolated during the course of this dissertation (Chapter III), except for the ATCC (American Type Culture Collection) strains and strain PWH3a which was provided by A. Chan (UBC). Species Strain Source Vibrio parahaemolyticus J00-5 Jericho Pier, BC Vibrio parahaemolyticus J00-6a Jericho Pier, BC Vibrio parahaemolyticus J00-7a Jericho Pier, BC Vibrio parahaemolyticus J00-9a Jericho Pier, BC Vibrio parahaemolyticus SOO-I Strait of Georgia, BC Vibrio parahaemolyticus S00-2 Strait of Georgia, BC Vibrio parahaemolyticus SOO-3 Strait of Georgia, BC Vibrio parahaemolyticus S00-5 Strait of Georgia, BC Vibrio parahaemolyticus S00-8 Strait of Georgia, BC Vibrio parahaemolyticus SO 1-20 Strait of Georgia, BC Vibrio parahaemolyticus SOI-31 Strait of Georgia, BC Vibrio alginolyticus ATCC 17749 ATCC Vibrio alginolyticus JOO-lOa Jericho Pier, BC Vibrio alginolyticus PWH3a Gulf of Mexico, TX Vibrio alginolyticus S99-45 Strait of Georgia, BC Vibrio alginolyticus S00-4 Strait of Georgia, BC Vibrio fluvialis S99-44 Strait of Georgia, BC Vibrio natriegens ATCC 14048 ATCC Vibrio vulnificus S99-48 Strait of Georgia, BC Vibrio vulnificus S01-38 Strait of Georgia, BC 15 2 .4 Results 2.4.1 Physical Parameters Water temperatures (Fig. 2.1, Table 2.2) within the oysters varied from ca. 12 to 14°C in summer and decreased to 4°C during winter. Salinity from the site ranged from a low of 20 psu in summer to near 30 psu during winter. 2.4.2 Bacterial and Viral Abundances With the exception of TPV in oysters (with salinity), bacterial abundances (Fig. 2.2A) were significantly correlated with temperature and salinity (Table 2.3), with highest abundances occurring in summer when temperatures were >10°C and salinities <26 psu. The highest bacterial abundance was in June when TPV within oysters was 7 x 104 cells cc"1, while in the O 1 water, TPV peaked in early July at 5.4 x 10 cells mL" . TPV were undetectable in oysters (<10 cells cc"1) and in the water (<1 cell mL"1) during winter. Vibrio parahaemolyticus in oysters followed a similar pattern to TPV, but peaked 3 weeks later at 2.3 x 103 cells cc"1 before reaching undetectable levels (<3 cells cc"1) by late September. The abundance of V. parahaemolyticus in oysters was correlated to TPV abundance in the water (r = 0.71, p < 0.05) and not in oysters (r = 0.48, p = 0.10), and the correlation was even stronger (r = 0.84, p < 0.05) if the 3 week shift between maximum abundances was incorporated. In contrast to bacteria, viruses in oysters were always detectable and there was no seasonal pattern to their abundance (Fig. 2.2B). Titers ranged from 5.0 x 103 to 1.1 x 105 viruses cc"1, with an average abundance of 3.3 x 104 viruses cc"1 (Table 2.2). Although viral abundance appeared to oscillate out of phase with oyster TPV during summer, the negative correlation between the two parameters was not significant (r = -0.24, p = 0.45). In fact, viral abundance was not correlated to any of the measured parameters (Table 2.3). Viruses were always undetectable (<10 mL"1) in the water column. 16 May Jun Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May Month (2000-2001) Figure 2.1. Physical data at the study site. Temperature (•) and salinity (O) during May 2000 to May 2001. 17 Table 2.2. The range of biological and physical parameters obtained from this study and others. If only one value was reported, that value is marked as the mean value. VpV: Vibrio parahaemolyticus Virus. Parameter Unit Min Mean Max Location Source Abundances - oyster Virus (VpVs) viruses cc"1 5.0 x 103 3.3 x 104 1.1 x 105 Coastal BC This study Vibrio spp. cells cc"1 <10 2.5 x 104 7.5 x 104 Coastal BC This study Vibrio parahaemolyticus cells cc"1 <3.0 243 2.3 x 103 Coastal BC This study Abundances - water Virus (VpVs) viruses L"1 <0.05 3.4 85 Coastal BC Chapter V <1.0 16 180 Coastal HI Paul etal., 1997 50 380 1.9 x 103 Coastal FL/HI Kellogg etal., 1995 Vibrio spp. cells mL"1 <1.0 710 5.4 x 103 Coastal BC This study Vibrio parahaemolyticus cells mL"1 <0.01 0.07 0.46 Chesapeake Bay Kaper etal., 1981 0.02 0.24 0.68 Coastal US DePaola etal., 1990 <0.01 2.7 550 Okayama, JP Chowdhury etal, 1990 <1.0 34 70 Pacific NW, US Kelly and Stroh, 1988 Physiology Bacterial Growth Ratea d 1 0b 0b 0.33° - US FDA, 1999 Oyster Grazing Rate d 1 — 0.05 - - Dame, 1996 Burst Size viruses cell"1 5.0 130 350 - Hidaka and Tokushige, 1978 Physical Temperature °C 4.0 9.4 14 Coastal BC This study Salinity psu 20 24 31 Coastal BC This study a Rates correspond to published observations at the minimum, mean, and maximum temperatures listed for this study. b These temperatures (4.0/9.4°C) are usually reported as bacteriostatic for V. parahaemolyticus. cThe reported rate of 0.33 d"1 was at a temperature of 15°C instead of the corresponding 14°C in this study. oo V 5 o o £ 3 o o c re TJ c 3 .Q re o 2 J -1 6 2, 5 o c re TJ c 3 re o 4 J Oyster T P V Water T P V 0 J Oyster Vp - i 1 1 1 1 1 1 r Jun Jul Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Month (2000-2001) Figure 2.2. Bacterial and viral abundances during May 2000 to May 2001. Closed symbols indicate abundances within oysters and open symbols indicate water-column abundances. (A) Bacterial abundances: water-column total presumptive Vibrio spp. (TPV, •), oyster total presumptive Vibrio spp. (TPV, •), oyster Vibrio parahaemolyticus (Vp, •). (B) Viral abundance in the oyster (A). Values are means of duplicate samples; error bars = range and are smaller than the width of the symbol if not visible. 19 Table 2.3. Pearson correlation analysis results of the physical and biological parameters measured in this study (from Figs. 2.1 and 2.2). Correlation values (r) are followed by p-values (two-tailed) and asterisks indicate correlations that are significant at p < 0.05. TPV: Total presumptive Vibrio spp.; Vp: Vibrio parahaemolyticus; VpV: Vibrio parahaemolyticus Virus. Water TPV Oyster TPV Oyster Vp Oyster VpV Oyster Temp. Water Salinity Water TPV 1 0.72* p < 0.05 0.71* p < 0.05 -0.05 p = 0.87 0.85* p < 0.05 -0.79* p < 0.05 Oyster TPV 1 0.48 p = 0.10 -0.24 p = 0.45 0.79* p < 0.05 -0.36 p = 0.25 Oyster Vp 1 -0.08 p = 0.82 0.58* p < 0.05 -0.66* p < 0.05 Oyster VpV 1 -0.22 p = 0.50 0.29 p = 0.36 Oyster Temp. 1 -0.72* p < 0.05 Water Salinity 1 2.4.3 Host Range The viruses were able to infect most strains of V. parahaemolyticus from the oysters and the sediment (62 and 74% of the pairings, respectively), but infected fewer strains from the water (30% of the pairings; Table 2.4). Collectively, the 13 viral strains infected 17 of 19 strains of V. parahaemolyticus, with each virus infecting a median of 11 strains (range 5 to 14). On average, each strain of V. parahaemolyticus was sensitive to ca. 7 of the 13 viruses (53% coverage of the total pairings) and ranged from two strains that were immune to all of the viruses to three strains that could be infected by 12 of the viruses. Ten of the viruses were not species-specific and infected a few (4 of 9) V. alginolyticus strains (the closest species to V. parahaemolyticus), one V. natriegens strain, and one of two V. vulnificus strains (Table 2.4). There was also a temporal trend in the patterns of infection (Fig. 2.3). The total number of bacterial strains infected by the viruses (i.e.: sensitivity) obtained when V. parahaemolyticus was present (summer) showed a significant decreasing trend (r2 = 0.71, p < 0.05) as the season progressed. The viruses infected from 12 to 18 (of 30) strains in June through August and decreased to 8 or less in September, the end of the period when V. parahaemolyticus was present, before rising to 15 and 10 (Table 2.4) for the two viruses isolated during the winter. 20 Table 2.4. Host range of VpVs isolated during summer 2000 and winter 2000/1 against co-occurring Vibrio parahaemolyticus (from 3 environments) and related Vibrio spp. Solid squares indicate lytic infections, whereas open squares indicate the absence of lysis. Strains with accompanying dates were isolated from Ladysmith Harbour during this study; other strains are from different dates and sources (see Table 2.1). "W": "Winter". "Summer" "yy" < < < < < < < < "T-t < T- t < < •T-t < T- t < •T-t Virus )V LH1 )V LH: )V LH: )V L» )V LH' )V LH' )V LHi )V LH? )V LHi )V LH( )V LH( )V LH' )V LH? P P o P a. P o P Jul-2 > > > > 00 CD Oo GO CD 00 00 <—i Host (Species and Strain) Date in-21 Jul-2 OQ i OQ i to ig-29 c OQ i to •a • T3 1 T3 1 :p-26 :p-26 O < i an-9 in-21 ig-29 :p-26 :p-26 ON an-9 V. parahaemolyticus V408-10 Jun-8 * >1 _< V. parahaemolyticus V416-7 Jun-21 " % *f .1 o V. parahaemolyticus V424-A3 Jul-6 V. parahaemolyticus V439-A1 Jul-27 111 lllill •K" •"Si n V. parahaemolyticus V465-A1 Aug-15 • j i V. parahaemolyticus V478-1 Aug-29 V. parahaemolyticus V497-B1 Sep-26 llilll V. parahaemolyticus C00-9a Sep-13 H I V. parahaemolyticus JOO-5 -Wat< V. parahaemolyticus J00-6a -Wat< V. parahaemolyticus J00-7a • - -'• ••• i V. parahaemolyticus J00-9a -V. parahaemolyticus S00-1 -V. parahaemolyticus SO 1-31 -V. parahaemolyticus S00-2 -C/3 n V. parahaemolyticus S00-3 - Piili :*<• IlliPf BP dim V. parahaemolyticus S00-5 - lliill lliill n a V. parahaemolyticus S00-8 - * Bill V. parahaemolyticus S01-20 -V. alginolyticus C00-6a Jul-27 PR HI V. alginolyticus V478-16 Aug-29 V. alginolyticus V478-17 Aug-29 O V. alginolyticus V497-C16 Sep-26 ET rt V. alginolyticus ATCC17749 -V. alginolyticus S99-45 -«" V. alginolyticus - 3 others -•3 V V.fluvialis S99-44 -V. natriegens ATCC14048 -V. vulnificus S99-48 -V. vulnificus S01-38 -21 Figure 2 . 3 . Total number of bacterial strains infected by each viral strain from oysters in summer (sensitivity). Viruses are organized in increasing isolation date and correspond to those in Table 2.4. 22 2.4.4 Plating Efficiencies The efficiencies (Fig. 2.4) with which summer strains of V. parahaemolyticus could recover infectious viruses were tested using a virus sample from summer and another from winter, when V. parahaemolyticus was below the detection limit. Bacterial strains from June and July recovered approximately half of the summer viral community, as compared to the control strain (El-80); whereas, for the winter viral community only one strain (V439-A1), one of the most permissive of the summer (Table 2.4), recovered a substantial portion (-51%) of the winter viral community relative to the control strain. Three summer strains of V. parahaemolyticus from June and July had very low plating efficiencies (-0.5-3%) while the 3 strains obtained in August and September did not recover any of the >5 x 104 VpVs in either sample. Among the 4 strains that were sensitive, the mean recovery was just 14% of the winter community compared to 48% of the summer community. 23 120 110 100 90 80 70 60 50 H 40 30 20 H 10 0 49 0.5 51 3.3 M L 3 8 0.6 54 m • Summer El Winter E1-80 V408-10 V416-7 V424-A3 V439-A1 V465-A1 V478-1 V497-B1 Control Jun-8 Jun-21 Jul-6 Jul-27 Aug-15 Aug-29 Sep-26 Host and Source Date Figure 2.4. Plating efficiency of summer versus winter oyster viral communities on strains of Vibrio parahaemolyticus from oysters in summer. Host strains and source dates correspond to those in Table 2.4. Efficiencies (%) are indicated on the figure for clarity and are relative to the control strain (El-80). Values are means of duplicate samples; error bars = range and are nearly 0 (or = 0) if not visible. 24 2.5 Discussion To my knowledge, this is the first study to follow the seasonal dynamics (inside, and outside, of oysters) of Vibrio spp. and the viruses which infect them. These data were combined with data on the host range of viral strains and the efficiency with which individual strains of Vibrio parahaemolyticus recovered infectious viruses from oyster and seawater samples. Together these data provide a comprehensive picture of changes in the host and virus communities over several months. Within oysters, seasonal changes of the abundances of Vibrio spp. and the host range of the viruses that infect them, along with the persistence of virus year-round, suggest two distinct regimes. There is a summer regime during which viral production in bacterial populations within oysters is presumed to occur, and a winter regime during which the virus population persists in the absence of detectable hosts. 2.5.1 Summer Viral Production within Oysters Two observations suggest that from June through August, when temperatures exceeded 10°C, conditions were favourable for production of viruses infecting Vibrio parahaemolyticus (VpVs). First, high abundances within oysters of potential host cells (total presumptive Vibrio and V. parahaemolyticus) and VpVs suggest high contact rates between viruses and hosts. Second, summer strains of Vibrio parahaemolyticus from oysters were susceptible to most of the summer VpV strains. An estimate of the effect of viral lysis on V. parahaemolyticus within oysters can be made based on approximations of burst size and viral turn-over time. This estimate can be used to infer whether V. parahaemolyticus could support VpV production. Although the burst size varies widely for V. parahaemolyticus (Table 2.2), a typical estimate for bacteria in the sea is 50 viruses cell"1 (Wommack and Colwell, 2000). Hence, to produce the average of 3.3 x 104 VpV cc"1 (Table 2.2) would require the lysis of approximately 660 cells. Many depuration and uptake studies have shown that bacteria and viruses are not accumulated by oysters, but that there may be low-level persistence, yet at abundances generally well below the levels observed in this study (reviewed in Richards, 1988). Oysters can only graze -5% of the bacterial size-fraction while the virus size-fraction should not be retained at all (Dame, 1996). This implies there is a constant exchange of bacteria and viruses between oysters and seawater. The few depuration studies that have looked at the rate of bacteriophage removal (Hoff and Beck, 1965; Hoff et al., 1966; Baross et al, 1974; Muniain-Mujika et al., 2002) observed complete (>99%) turnover of 25 large abundances in an average of -2 d, with some persistence of a trace amount for longer periods. Other studies that have found slower depuration rates, or persistence of larger titers, have examined RNA phage and eukaryotic RNA viruses (reviewed in Richards, 1988) that may have particularly longer residence times due to their smaller size. Therefore, assuming the vibriophage population turned over every two days, ca. 136% of the average V. parahaemolyticus standing stock during summer (243 cells cc"1, Table 2.2) would be lysed daily by viruses in order to sustain the measured VpV titers within oysters. This represents over 5 times the daily bacterial production at the maximal growth rate of 0.33 d"1 that has generally been reported for V. parahaemolyticus within live shellstock at the maximal summer temperature (Table 2.2). Only when virus titer was lowest (5.0 x 103 cc"1) and bacterial abundance highest (2.3 x 103 cc"1) could the estimated mortality of 55% of daily production be near reasonable. Although V. parahaemolyticus populations can grow at rates of up to 36 d"1 for 90 min when oysters are uncovered at low tides, these spikes in bacteria are washed out -1-2 h after the tides re-submerge the oysters (Buenaventura et al, 2002). Even if bursts of phage production were to occur at these times, they would be washed out as well and there is a limited window in the summer season (clear days, high air temperature) during which this would be applicable. Although these figures give us a sense of the capacity for VpV production within oysters, they must be interpreted with caution. Rates of depuration and retention observed by others are highly variable, presumably due to the myriad of physical conditions (e.g.: types of systems, temperatures, etc.), mollusc species, and virus types that have been used in these studies. Furthermore, growth estimates for V. parahaemolyticus from the Pacific Northwest have not been done and may show better cold-adaptation, and hence higher growth rates at the observed temperatures, than more southern strains. Consequently, if the depuration rates of VpVs were -2-fold slower and the growth rates of V. parahaemolyticus -3-fold higher (nearer 1 d"1), production of VpVs could be reasonably supported without the requirement for additional host species. Given this variability and the fact that other Vibrio spp. can serve as hosts (Table 2.4) for VpVs, it is reasonable to conclude that viral production during summer may be sustainable within oysters because of the presence of multiple host species. This requirement is, however, more applicable during colder periods. Nonetheless, the results emphasize that viruses were significant agents of bacterial mortality within these oysters. 26 2.5.2 Summer Viral Production in the Water Column and Sediment In contrast to the findings within oysters, there appears to be little production of VpVs within the water column and sediment. Viruses infecting V. parahaemolyticus were undetectable (<10 mL"1) year-round in water surrounding the oysters, although low abundances of VpVs in seawater have been detected in the coastal waters of BC and elsewhere (Table 2.2). This suggests that seawater is not a source of VpV but can serve as a medium of transmission of VpVs among oysters. The abundances of V. parahaemolyticus typically found in seawater are also low (Table 2.2) indicating that contact rates between VpVs and their hosts, and hence rates of lytic virus production, would also be low. In sediments from coastal BC, VpVs have not been detected (Chapter V); however, V. parahaemolyticus strains were readily found and very susceptible to infection by VpVs, indicating the potential for VpV production. It is possible that the V. parahaemolyticus strains obtained from the sediments were from the same population isolated from the oysters, given that both occurred in the same inlet where oyster aquaculture is extensive. Therefore, the evidence as a whole suggests that the high abundance of VpVs in oysters was produced by bacteria within the oysters. It is unlikely that the VpVs were concentrated from the surrounding seawater given that they would have to be preferentially retained over the much higher abundances of morphologically similar viruses. The schematic in Figure 2.5 gives a general representation of the host-virus dynamics in summer (A panel), in which V. parahaemolyticus and other bacteria serve as hosts for viral production. Given the high production rates of V. parahaemolyticus and VpVs within oysters, which are continuously shed into the environment, the oysters serve as the major environmental source of V. parahaemolyticus and VpVs; and serve as seed populations for other locations (sediment, zooplankton, molluscs) and seasons, transmitted via the water column. 27 A: Summer Figure 2.5. Schematic representation of Vibrio parahaemolyticus-VpV dynamics in oysters. The oysters are represented by the boxes and the sediments by the gray zone. (A) During summer, viruses are produced by growing V. parahaemolyticus cells (closed circles) and by contributions of other species (probably related Vibrio spp.; open circles). There is continual exchange of viruses and bacteria with the water column (double arrows). (B) During winter, the virus population is persistent, whereas cells of V. parahaemolyticus are only known to be associated with zooplankton and in sediments. Virus production therefore takes place in host cells of different species and/or cells that have altered culturability (dashed circles). 28 2.5.3 The "Winter Conundrum" - Viral Persistence Frequently, there are strong seasonal variations in viral abundance which mirror those of total bacterial abundance (Jiang and Paul, 1994) or those of specific hosts (Waterbury and Valois, 1993). Therefore, one would predict that the winter disappearance of V. parahaemolyticus would be mirrored by the disappearance of VpVs. However, these data show that vibriophages persist in oysters when host cells are undetectable. Similar results have been found for the Gulf of Mexico and the Pacific Northwest (Baross et al., 1978b; DePaola et al., 1998). The persistence of VpVs in winter may result from viral production that is supported by bacteria other than V. parahaemolyticus (Table 2.4), or by host populations that persist but which are not detectable during the winter. The following hypotheses can be formulated to explain the persistence of the viral population over winter when Vibrio spp. are undetectable: 1) the virus population persists in the absence of viral production; 2) viral production occurs within hosts in the viable but nonculturable (VBNC) state; 3) viral production occurs within non-Vibrio spp. The first hypothesis requires that there is minimal decay or dilution of viruses within oysters, and assumes the absence of VpV production given that V. parahaemolyticus and related Vibrio spp. are undetectable during winter (Fig. 2.2; Kaneko and Colwell, 1978; Chowdhury et al, 1990; Zaccone et al, 1992). Marine bacteriophage can be stored for several months with minimal decay in natural seawater at 4°C; hence it is conceivable that the viruses could persist during the winter. Although the pumping rates of molluscs decrease during colder periods (Dame, 1996), continuous circulation is still required for feeding and respiration. Therefore, without production VpV abundances within the oysters should decrease until they are equal (undetectable) with the surrounding seawater (Baross et al, 1974; Richards, 1988). The second hypothesis allows for viral production to occur during winter in the appropriate host, but within cells that were not quantified. Vibrio parahaemolyticus over-winters in sediments and zooplankton, and is generally undetectable in molluscs and the water column during cold months (Kaneko and Colwell 1973; Kaneko and Colwell 1978; Venkateswaran et al, 1990; Grimes, 1991; Fig. 2.5B). A recent study found that V. parahaemolyticus and V. vulnificus persisted in an estuary during winter (Fukushima and Seki, 2004), but at levels (-10" MPN mL"1) below those expected to support significant viral production. Vibrio parahaemolyticus (and related Vibrio spp.) may enter into the viable but nonculturable (VBNC) state within -2 months at 4°C (Chowdhury et al, 1990; Jiang and Chai, 1996). However, it is not known if viral replication can occur in these cells. It is possible that a state of unculturability 29 ("cold adaptation") exists that would allow infection, or that "summer strains" (mesophiles) of V. parahaemolyticus may be displaced by "winter strains" (psychrophiles) that are not culturable on the standard nutrient-rich medium at 25°C. The lack of reported illnesses in winter from mollusc consumption (Canadian Food Inspection Agency, unpubl.) suggests that V. parahaemolyticus is not present. Alternatively, pathogenic strains (tdh+) may be more adapted to warmer temperatures and be out-competed by non-pathogenic (tdh) strains during colder months (Pace and Chai, 1989; Jiang and Chai, 1996). The final hypothesis is that VpVs infect other species that sustain viral production over the winter. Vibrio parahaemolyticus comprised <14% of the cells belonging to Vibrio spp. and presumably a much smaller portion of the total bacteria within oysters. Although there are very few described Vibrio spp. (V. logei, V. marinus) that are psychrotolerant or psychrophilic (Baumann and Shubert, 1984), unnamed Vibrio spp. can be readily isolated, and at times dominate, among 4°C bacteria (Urakawa et al, 1999; Radjasa et al, 2001). The high titers of VpV (104-105 cc"1) combined with a host range that extends to other species (Table 2.4) provides strong evidence that high abundances of VpVs persist within oysters during winter because of viral production sustained by other species (Fig. 2.5B). This is consistent with the calculations above that indicated production of VpV in summer was also supported by species other than V. parahaemolyticus. In winter, the virus community is less able to infect summer host strains (Fig. 2.3), as is shown by the plating-efficiency data (Fig. 2.4). These data clearly demonstrate that the viral community composition has changed. There are other examples of seasonal changes in viral communities that infect specific hosts. For example, viruses infecting Synechococcus spp. have higher resistance to UV radiation in summer than in winter (Garza and Suttle, 1998). Therefore, winter VpV production was not supported by summer bacterial strains. Ultimately, the system is probably a dynamic mixture of altered host culturability and viral exploitation of cross-species infection. Through the examination of seasonal changes in the abundances of bacteria and the viruses that infect them, this study has shown that bacteria within oysters are likely the primary environmental source of viruses infecting V. parahaemolyticus. As well, seasonal shifts in patterns of host range provide strong evidence that a significant shift in the virus community occurs during winter, and although these viruses can infect many strains of V. parahaemolyticus, the persistence of these viruses in oysters during winter is likely supported by other hosts, possibly including closely related, but psychrophilic, bacterial strains. 30 Chapter III: Impact of Selection on the Distribution and Diversity of Vibrio spp. in Coastal Environments 31 3.1 Summary This study examined the distribution, phage-typing patterns, and genetic diversity of Vibrio spp. in the coastal waters of British Columbia in order to examine the impact of selection within different environments on hosts of marine phage. Abundances of Vibrio spp. ranged from 1.5 to 346 mL"1 throughout the water column (1-291 m). Abundances at the water-sediment interface were much higher (up to ~2.8 x 104 Vibrio spp. cc"1), and decreased with sediment depth (down to 30 cm). The widespread presence of Vibrio spp. in the areas investigated and their phage-typing patterns showed that they could support vibriophage production. Not surprisingly, geographic distribution was a poor determinant of bacterial diversity. Environment of isolation (water, sediment, or oyster) however, had a much more profound effect on genetic diversity (as revealed by ERIC-PCR), and on phage-typing patterns to an even greater extent. The patterns of susceptibility of Vibrio spp. isolates to different phage (from the water column and oysters) demonstrated that sources of high abundance (sediment and oysters) generally produced V. parahaemolyticus strains with high susceptibility to viral infection; and the opposite was true for the water column which produced highly resistant strains. These results show that there is segregation of bacterial host strains in different environments, under differing selection pressures, which will have an impact on in situ phage production. 32 3.2 Introduction Members of the Vibrionaceae are halophilic, gram-negative bacteria found in many aquatic environments (reviewed in Thompson et al, 2004a). They can compose up to 30%, but more typically 0.1-10% of the bacterioplankton as determined by culturing on solid media (Kaneko and Colwell, 1973; Zaccone et al, 1992) or, more recently, by analysis of 16S rRNA (Thompson et al, 2004b). They have wide tolerances for physical and chemical conditions (Baumann and Shubert, 1984), and can be found in high abundances in sediments, mollusc tissues, and zooplankton (Kaneko and Colwell, 1978; Arias et al, 1999; Pfeffer et al, 2003). Vibrio spp. have often been used as model prokaryotes in many culture-based and in situ experiments, including as hosts in studies of marine phage ecology (Suttle and Chen, 1992; Proctor et al, 1993; Jiang and Paul, 1996; Moebus, 1996; Weinbauer etal, 1997). Vibrio parahaemolyticus is typically found in estuarine (Fukushima and Seki, 2004) and coastal temperate (Chowdhury et al, 1990; Alam et al, 2003) and sub-tropical waters (Cavallo and Stabili, 2004), and can cause gastroenteritis when contaminated raw shellfish are consumed (Grimes, 1991). Highest abundances generally occur in sediments and mollusc tissues (10 and 104 CFU g"1, respectively) when temperatures are >15°C and salinities <30 psu (Baross et al, 1978b; Kaneko and Colwell, 1978; Chowdhury et al, 1990; DePaola et al, 2003). The diversity of V. parahaemolyticus strains has been studied; however these investigations are generally within the context of food protection (Venkateswaran et al, 1998) and clinical epidemiology (Shirai et al, 1990). Much less is known about the genetics of environmental strains or the susceptibility of V. parahaemolyticus to viral infection. The purpose of this study was to determine the extent of Vibrio spp. populations in several environments along the coast of British Columbia (BC) and to determine whether there were geographic trends in their genetic diversity and phage-typing patterns (phage susceptibility). As a larger goal, there is interest in seeing what these diversity trends mean in terms of these bacteria acting as hosts for vibriophage populations. These hosts are being used, within this dissertation, as a model system to assess whether there is segregation of heterotrophic bacterial strains, under differing selection pressures in different marine environments, which will have an impact on in situ phage production. 33 3.3 Materials and Methods 3.3.1 Sample Collection and Processing Water and sediment samples were collected from coastal BC (Fig. 3.1 and Table 3.1) from 1999 to 2003. The samples were collected during annual research cruises in July and August, except for the Jericho Pier samples, which were collected from June to September 2000. Water samples were collected using Go-Flo bottles mounted onto a rosette, from an onboard seawater intake (depth = 2 m), or by surface sampling with a bucket (Jericho Pier only). During the cruise in 2002, three transects comprising 21 stations with ~1 km spacing were conducted within the Malaspina system (Fig. 3.2). During 2003, a transect was conducted within the Fraser River plume (Fig. 3.3) comprising 9 stations, each separated by -5.5 km (3" of latitude). Sediment core samples were collected using a triple-barreled gravity corer (Rigosha and Co.) and pore water was extracted as described previously (Lawrence et al., 2002). Briefly, the sediment-water interfaces were sampled using serological pipettes and deeper samples were sliced from the extruded cores. Twenty cubic centimetres of each sample were mixed with an equal volume of PBS (Sambrook et al, 1989) in a 50 mL polypropylene tube and agitated on a tabletop shaker for 30 min. at 150 rpm. Samples were then centrifuged at 4000 x g for 60 s to separate coarse sediment materials from pore waters. The pore waters were sub-sampled for Vibrio spp. counts (see below), and the volumes of extruded water and remaining sediment were recorded. Table 3.1. Total Presumptive Vibrio spp. (TPV) samples from coastal BC and corresponding ranges of measured values. Temp.: temperature; Sed.: sediment. Number of Depth Temp. Salinity CFU Locations Samples Water(m) Sed. (cm) (°C) (psu) mL'orcc 1 Surface water 27a 75 1-10 - 7-21 11-33 1.5-346 Deep water 11 17 11-291 - 7-13 28-33 1.0-63 Surface sediment 14 19 23-265b 0-lc - - 1.0-28300 Deep sediment 9 28 34-265b 1-30 - - 0.7-8460 Total: 139 a Excluding the small-scale station locations for the Malaspina and Fraser River plume transects. b These values refer to the depth of the water column above the sediments. 0 This value represents the water-sediment interface, going into the sediment to a depth of 1 cm. 34 3.3.2 Physical Properties A CTD (conductivity/temperature/depth; Seabird Electronics), outfitted with a fluorometer (WET Labs), was mounted on the rosette and used to measure in situ profiles of temperature, salinity, and chlorophyll fluorescence (relative units). Temperature and salinity of the samples taken along the transects and at Jericho Pier were taken using a thermometer and a hand-held refractometer (Fisher Scientific). 3.3.3 Growth Media and Conditions All bacteria were suspended in ultrafiltered (30 kDa cut-off, Millipore; Suttle et al, 1991), sterilized natural seawater diluted to 15 psu with reverse-osmosis water (Millipore). The purification and maintenance medium was Marine Luria-Bertani broth (MLB; 0.5 g L"1 each of casamino acids, peptone, and yeast extract, 0.3% v/v glycerol, in 15 psu ultrafiltrate base). This broth was amended with 1% w/v agar for solid plates. All liquid cultures, when needed for assays, were grown at 30°C with agitation (150-200 rpm). Plaque assay and bacterial stock plates were grown at room temperature (~25°C). 3.3.4 Total Presumptive Vibrio spp. (TPV) Enumeration and Isolation To obtain estimates of mesophilic TPV abundance, water and pore-water sub-samples were serially diluted in duplicate 10 mL (final volume) of sterile seawater (as mentioned above). The dilutions were filtered onto 0.45 urn pore-size cellulose membranes (47 mm HAWP, Millipore) which were laid, two per plate, onto TCBS agar (BD Gibco). The plates were incubated at 25°C for -24 h. Colonies were purified by serially re-streaking 3 times onto MLB plates. Pure strains were then identified biochemically (API 20 E®, bioMerieux). 3.3.5 Phage-Typing Analysis Bacteria isolated during this study (Table 3.2) were phage-typed using viruses isolated from oysters and the water column of the Strait of Georgia. Reference strains (e.g. ATCC, BCCDC), as well as bacteria isolated from Strait-of-Georgia oysters, were also included in the phage-typing to serve as comparison strains. The 29 viral strains are described in detail elsewhere (Chapters II and V). Ten to twenty microlitres of amplified virus stocks were spotted into 0.6% agar overlay lawns of each bacterium on MLB plates and monitored for zones of clearing for 1-7 d. The results of 928 crosses (32 Vibrio spp. x 29 viruses) were scored as presence (zone of lysis = "1") or absence (no lysis = "0") data. A distance matrix was then calculated using the Dice 35 similarity coefficient (2nab / [na + nt,]; Dice, 1945), which is basically the ratio of things in common, out of the total, between each pair-wise combination of bacteria. Twice the number of shared elements (2nat,; in this case shared "Os" and "Is") between any two bacteria is divided by the total number of elements in both bacteria (na + nt,; total number of "Os" and "Is"), resulting in a value between 0 and 1. This distance matrix was then used to create a UPGMA tree using the NEIGHBOR function in PHYLIP v3.5 (http://evolution.genetics.washington.edu/phylip.html). 3.3.6 Induction ofLysogens Exponentially-growing bacterial strains (media and conditions above) were treated with 1 pg mL"1 (final concentration) of mitomycin C (Sigma) and incubated for 3 days. Aliquots were then centrifuged at 15,000 x g for 3 min to pellet cell debris and the supernatants treated with DNase I (125 Kunitz units; Sigma) and RNase A (11 Kunitz units; Sigma) for 30 min at 37°C to remove any residual naked nucleic acids. Finally, supernatants were processed for epifluorescence virus counts according to the modified procedure of Noble and Fuhrman (1998) presented in Wen et al. (2004). Briefly, supernatants were filtered through 0.02 pm Anodiscs (Whatman) and the filters were then placed face-up onto a drop of SYBR Green I nucleic acid stain (Molecular Probes). The filters were incubated in the dark for 15 min then mounted onto slides using 50% glycerol-50% PBS (Sambrook et al, 1989) amended with 0.1% p-phenylenediamine (Sigma). Filters were observed at lOOOx magnification (Olympus AX70) under wide-blue excitation (450-480 nm, 515 nm cut-off). 3.3.7 Genetic Fingerprinting and Analysis Genetic fingerprinting was carried out using the Enterobacterial Repetitive-Element Intergenic Consensus Sequence PCR (ERIC-PCR; Versalovic et al, 1991). The 50 uL ERIC-PCR reactions consisted of the following reagents (Invitrogen Life Technologies): Taq DNA polymerase buffer, 3 mM MgCl2, 0.4 mM of each deoxyribonucleoside triphosphate, 2 uM of each of the ERIC1R and ERIC2 primers, and 1 U of PLATINUM Taq DNA polymerase. Negative controls contained all reagents except template DNA. PCR was carried out with the following cycle parameters: initial denaturation at 95°C for 1.5 min.; followed by 35 cycles of denaturation at 95°C for 1 min., annealing at 52°C for 1 min., extension at 72°C for 2 min.; and a final extension at 72°C for 10 min. PCR products (10 p:L each) and 100 bp ladders (Invitrogen Life Technologies) were electrophoresed in 1.5% agarose in 0.5X TBE buffer (45 mM Tris-borate, 1 mM EDTA [pH 8.0]) at 110 V for 3 h. Gels were stained with ethidium bromide and 36 captured with an Alphalmager 3400 gel documentation system (Alpha Innotech Corporation). Digital gel images were then cropped and enhanced (brightness/contrast) using Adobe Photoshop 5.0 LE. Fingerprints were compared quantitatively using GelCompar II (Applied Maths). To quantify relationships among strains, UPGMA trees were constructed using a Pearson correlation method that compares the densitometry profiles of each lane to one another. For a more in-depth explanation of the GelCompar analysis, consult Ferrari and Hollibaugh (1999). 3.3.8 Statistical Analysis Summary statistics, tests for normality (Shapiro-Wilk), and correlation analyses (Pearson and Spearman) were carried out with the PAST program (http://folk.uio.no/ohammer/past/). Median values, and not mean values, are reported because the datasets had non-normal distributions. Bacterial abundances in the sediment were expressed per cubic centimetre of original sediment volume after taking into account extraction dilution. 37 Table 3.2. Bacterial strains (isolated in this study and others) used to determine host susceptibility and genetic fingerprinting patterns. Malaspina strains are additionally marked with the more accurate locations (LA/MA/OK) corresponding to Fig. 3.2. ATCC: American Type Culture Collection; BCCDC: British Columbia Centre for Disease Control. Species Strain(s) Environment Location Source Vibrio parahaemolyticus ATCC17802 - - ATCC Vibrio parahaemolyticus C00-9a Water Ladysmith Harbour, BC Chapter II Vibrio parahaemolyticus El-80 and E5-19 - - BCCDC • Vibrio parahaemolyticus J00-5 to J00-9a Water Jericho Pier, BC This study Vibrio parahaemolyticus S00-1 Water Malaspina (LA), BC . This study Vibrio parahaemolyticus S00-2/3/5 Sediment Malaspina (LA), BC This study Vibrio parahaemolyticus S00-8 Sediment Malaspina (MA), BC This study Vibrio parahaemolyticus SO 1-20 Sediment Malaspina (OK), BC This study Vibrio parahaemolyticus S01-31 Water Nanoose Bay, BC This study Vibrio parahaemolyticus V408-10to V497-BI Oyster Ladysmith Harbour. BC Chapter 11 Vibrio alginolyticus ATCC 17749 1 • : i l l l l f i l : : ; l j l t l I - i l l l l ATCC Vibrio alginolyticus C00-6a Water Ladysmith Harbour, BC Chapter II Vibrio alginolyticus JOO-lOa Water Jericho Pier, BC This study Vibrio alginolyticus PWH3a Water Gulf of Mexico, TX Suttle and Chen, 1992 Vibrio alginolyticus S99-45 Water Pendrell Sound, BC This study Vibrio alginolyticus. S00-4 Sediment Malaspina (LA), BC This study Vibrio alginolyticus V478-16to V497-C16 Oyster Ladysmith Harbour, BC Chapter II Vibrio fluvialis S99-44 Sediment Malaspina (OK), BC This study Vibrio natriegens ATCC 14048 - - ATCC Vibrio vulnificus S99-48 Water Pendrell Sound, BC This study Vibrio vulnificus S01-38 Sediment Saltspring Island, BC This study 0 0 3.4 Results and Discussion 3.4.1 Horizontal and Vertical Distribution Total presumptive Vibrio spp. (TPV) in surface waters (upper 10 m) varied between 1.5 to 346 CFU mL"1 (median = 48 CFU mL"1; Table 3.1). Vibrio spp. were present at all sampled temperatures (7-21°C) and salinities (11-33 psu), but were not correlated to these parameters. However, abundances were generally higher (>50 CFU mL"1) near locations of active shellfish and salmon aquaculture (Fig. 3.1: AP, LI, MI, QI, SP, TC; Fig. 3.2) and the Fraser River plume (Fig. 3.1: JP; Fig. 3.3); and lower in more "pristine" locations, such as remote inlets like Pendrell Sound (Fig. 3.1: PS), and in more open-water areas, such as the Strait of Georgia and Queen Charlotte Strait (Fig. 3.1: QCS, SOG). There is high variability in the reported abundances of Vibrio spp. from coastal surface waters, ranging from <1 to a maximum of 3 x 105 CFU mL"1 off the coast of Italy (Venkateswaran et al, 1989b; Venkateswaran et al, 1990; Barbieri et al, 1999; Maugeri et al, 2004). More typically, mean abundances are 1-100 CFU mL"1 (Kaneko and Colwell, 1973; Venkateswaran et al, 1989a; Oliver et al, 1982; Hervio-Heath et al, 2002; Pfeffer et al, 2003; Cavallo and Stabili, 2004), corresponding to the range of values observed in this study. Spatial variability among surface samples was examined in the three inlets comprising the Malaspina Inlet system (Fig. 3.2). Salinity differences among the 21 stations sampled were only slightly different (Fig. 3.2A), ranging from ca. 29 to 31 psu. Generally, the TPV abundance was similar to the median (148 CFU mL"1; Fig. 3.2B), except for one station in Okeeover Inlet where it was ca. 2-fold higher. The increase was coincident with a slightly higher temperature compared to adjacent stations (Fig. 3.2C). However, temperature was probably not the main cause as similar temperature changes at other stations (LA #3-7) did not correspond to a change in TPV abundance. Additionally, colder (12-13°C) water penetrating into Malaspina Inlet (MA #1-5) from Desolation Sound was not associated with a decrease in TPV abundance. 39 126°W 124°W Figure 3.1. Geographic distribution of samples. Locations sampled for total Vibrio spp. are indicated by closed circles for water column only (•), or open circles for both water and sediment ( o ) . For reference, Jericho Pier (JP) is located in Vancouver. AP: Arrow Pass; DS: Desolation Sound; HO: Hotham Sound; HS: Howe Sound; JS: Johnstone Strait; LH: Ladysmith Harbour; LI: Larsen Island; MI: Malaspina Inlet; NB: Nanoose Bay; NI: Narrows Inlet; PS: Pendrell Sound; QCS: Queen Charlotte Strait; QI: Quadra Island; SA: Salmon Inlet; SN: Saanich Inlet; SOG: Strait of Georgia; SP: Saltspring Island; TA: Teakerne Arm; TC: Tribune Channel. 40 A : Geography & Salinity B : T P V Coun t s 50°N Figure 3.2. Malaspina system short-scale Vibrio spp. distribution. A: Locations of the ~1 km-spaced stations (•) along the three transects (MA, LA, and OK) with the corresponding salinity values. B: Distribution of TPV counts (means of duplicates) along the transects. The median value (148 CFU mL"1) is represented by the horizontal line as indicated. Typically, the range of duplicate determinations was within 25% of the median except for stations OK2 and OK3, which were 28 and 235% over the median, respectively. C: Temperature distribution along the transects. Temperature values are discreet and not ranges, therefore most of the inlet water was 16-18°C except for some 12-13°C water in MA. Note that, for B and C, station numbers begin outwards (three #1 stations) and increase towards the centre of the system (the three #7 stations in the middle). 41 The abundance of Vibrio spp. in the surface of the Fraser River plume was also investigated. The Fraser River has a plume that can cover a significant portion of the southern Strait of Georgia, affecting biological and nutrient dynamics (Harrison et al, 1983). The abundance of TPV decreased sharply from >100 to <50 mL"1 from Stations 1 to 4, as the salinity decreased from 22 to <14 psu. Similarly, from Stations 5 to 7, TPV decreased 10-fold from 70 mL"1 to 7 mL"1 with the decrease in salinity from 18 to -12 psu (Fig. 3.3). The presence of Vibrio spp. in estuarine waters is common, although abundances typically decrease along with salinity (O'Neill et al, 1992; Barbieri et al, 1999; Fukushima and Seki, 2004). As with the Malaspina transect, temperature was not related to TPV abundance. Vibrio spp. also occurred (median = 7.0 CFU mL"1) from the surface mixed layer to the maximum depth sampled of 291 m (Table 3.1). Although the relative abundances of TPV varied greatly, in most profiles (e.g. Fig 3.4, SOG) the abundance of TPV increased from the surface to peak at a sub-surface maximum, then decreased with depth, although not necessarily linearly (as in this example). Very few water-column profiles exist of Vibrio spp. below ca. 2 m depth, but those that do have shown 0.1-2 CFU mL"1 down to 180 m (Kaneko and Colwell, 1974; Venkateswaran et al, 1989b). Although marine bacterioplankton abundance and production generally correspond to the greatest phytoplankton biomass (chlorophyll a maxima - Andrade et al, 2004; Liu et al, 2004), this was not the case for Vibrio spp. The maxima in TPV were located below the chlorophyll maxima just as often as the two corresponded. The only unique, pattern was from Saanich Inlet (SN; Fig. 3.1 and Fig. 3.4), which has an intermittently anoxic bottom layer (Anderson and Devol, 1973). Here, TPV abundance increased with depth, with the bottom sample (210 m) being within the anoxic layer and having -12 times the surface abundance. All Vibrio spp. are facultative anaerobes (Baumann and Shubert, 1984), and therefore can thrive in these waters. 42 Figure 3.3. Distribution of Vibrio spp. in the Fraser River plume. A: Locations of the 3 minutes of latitude-spaced transect stations (•). Arrows indicate the general surface flow of the Fraser River plume. B: Distribution of Vibrio spp. CFU (bars), temperature (•), and salinity (A). Error bars (for Vibrio spp.) indicate the range of duplicate samples and are smaller than the width of the symbol if not visible. 43 Figure 3.4. Dis t r ibut ion of Vibrio spp. wi th depth. Depth profile of a typical station (Strait of Georgia, SOG; •) compared to the Saanich Inlet station (SN; •) where the bottom sample (grey star) was in the anoxic layer. Error bars indicate the range of duplicate samples and are smaller than the width of the symbol if not visible. 44 Vibrio spp. were up to -80 times more abundant in surface sediment samples (median = 231 CFU cc"1) than in the water column (Table 3.1). As for the water column, abundances were typically higher near aquaculture sites (Fig. 3.1: MI, QI, TC) and lower in more pristine areas (e.g. Pendrell Sound), although this was not true for all locations (LH, NB, and SP have aquaculture sites, yet were below the median). There was not a significant correlation between the abundances of Vibrio spp. in the sediments and the surrounding waters. Vibrio spp. were found up to 30 cm into the sediment, but were absent in the two deeper samples (35 and 40 cm). Abundances in the overlying water were typically less than at the water-sediment interface (e.g. Fig. 3.5), except for one location where abundances were near the detection limits. Similarly, in 9 cores TPV were higher at the water-sediment interface than deeper in the cores, while in two cores the highest values were just below the interface (at 1.5 and 5 cm). Typically, by 5 cm the TPV abundances were less than in the overlaying water. Other studies that investigated water-sediment interfaces or used grab samples of the top few centimetres found the abundances of Vibrio spp. to be highly variable (<1 to >105 CFU g"1), depending on season and geography (Kaneko and Colwell, 1974; Venkateswaran et al, 1989b; Venkateswaran et al, 1990; Pfeffer et al, 2003). 45 10 Vibrio spp. C F U cc" 1 100 1000 10000 Figure 3.5. Distribution of Vibrio spp. within sediment cores. Depth profile of a typical sediment core (Malaspina Inlet, MI; core depth 39 m) along with the corresponding value from the overlying water (W; 6.5 m). Error bars indicate the range of duplicate samples and are smaller than the width of the symbol if not visible. 46 3.4.2 Phenotypic and Genotypic Diversity Phage-typing patterns were used to infer the phenotypic diversity (Fig. 3.6) of Vibrio spp. isolated from the water column and sediment during this study. These susceptibility patterns were compared to those of strains from Strait-of-Georgia oysters, as well as reference strains (e.g. ATCC; Table 3.2). Based on the susceptibility patterns to 29 virus isolates, there were two major host-range clades (HR1 and HR2) separated by approximately 50% dissimilarity. The HR1 clade contains only V. parahaemolyticus strains and is composed of all of the sediment strains plus most (5/7) of the strains from oysters. Members of this clade group together since they were highly susceptible to viral infection. There is high divergence within the clade with all but one grouping of strains <80% similar. The HR2 clade contains a mixture of strains from V. alginolyticus, V. fluvialis, V. natriegens, V. parahaemolyticus, and V. vulnificus, although most of the clade "space" is occupied by V. parahaemolyticus and V. alginolyticus. All of the water-column strains of V. parahaemolyticus fell within this clade although they are interspersed among strains of V. alginolyticus. No sub-groups attributable to geographic origin of the strains, or other factors, were discernable because of the low divergence within this clade (nearly all one super-group). This low divergence results from the high resistance to viral infection shared amongst the 22 strains (69% of total) in this large clade (HR2). However, one sub-group consists of strains which were resistant to all nearly all viruses (centered near the "All neg." group). Vibrio fluvialis, V. natriegens, V. vulnificus, three V. alginolyticus strains (including the ATCC reference strain), and a water-column strain (J00-5) of V. parahaemolyticus all fall within this cluster. 47 0.1 Vp S01-20 Vp V408-10 Vp V465-A1 Vp S00-5 Vp S00-3 Vp V416-7 Vp V439-A1 Vp S00-8 Vp V424-A3 Vp S00-2 Va PWH3a Vp J00-6a Vp S00-1 Va S99-45 Vp V478-1 Va C00-6a Va V478-16 Va V478-17 Vp J00-9a Vn ATCC Vp J00-5 All neg. Va ATCC Vv S99-48 Vp C00-9a Vp J00-7a Vp V497-B1 Vp S01-31 HR1 HR2 Figure 3.6. Vibrio spp. phenotypic diversity. UPGMA tree based on the susceptibility of isolates of Vibrio spp. to infection by viruses. Similarities were calculated from 928 crosses (32 Vibrio spp. x 29 viruses). Two host-range clades (HR1 and HR2) are indicated. Vibrio parahaemolyticus strains are identified as follows: black shaded (with white text) are from sediments, grey shaded are from oysters, and within boxes are from the water column. Strain designations are listed in Table 3.2. The "All neg." group contains 5 strains which were not infected by any of the viruses: V alginolyticus J00-10a/S00-4/V497-C16, V. fluvialis S99-44, and V. vulnificus SOI-38. Va: V. alginolyticus; Vn: V. natriegens; Vp: V. parahaemolyticus; Vv: V. vulnificus. The scale bar represents a dissimilarity of 10%, corresponding to a difference of 5.8 infection/non-infection pairs between strains. 48 To determine whether the phage-typing patterns reflected genotypic diversity, ERIC-PCR was used to genetically fingerprint the strains (Fig. 3.7). Analysis of these patterns (Fig. 3.8) showed a relatively clear separation between V. parahaemolyticus (Vp clade) and the other Vibrio spp., suggesting that the V. parahaemolyticus strains SO 1-20 and SO 1-31 that cluster within the V. alginolyticus clade (Va) were misidentified biochemically. Vibrio alginolyticus and V. parahaemolyticus are phenotypically closely related and can easily be misidentified using biochemical tests on marine isolates (Robert-Pillot et al., 2002). The more distant relationship of V. vulnificus was also reflected in the ERIC-PCR similarities (bottom outgroup). It is noteworthy that within the V. parahaemolyticus clade of the tree (Vp), all of the "reference" strains (ATCC type-strain and the strains El-80 and E5-19; see Fig. 3.7) grouped within a separate sub-group (VpR) from the BC isolates (all other Vp). Embedded within the V. parahaemolyticus clade is a sub-group which contains 5 of the 7 strains from oysters (VpO). Interestingly, the two oyster strains missing from this sub-group (V478-1 and V497-B1) are the same two that did not group with the oyster/sediment cluster in the phenotypic tree (in HR2, not HR1; Fig. 3.6). Unlike the phage-typing tree, however, the sediment strains are not tightly grouped with the oyster strains genetically. The strains isolated from the water column and sediment are scattered throughout a few clusters consisting of two or three strains, and are not grouped with respect to geography nor environment (i.e.: sediment strains not all grouped together). Overall, only strains J00-6a and S00-1, and possibly V416-7 and V439-A1 (given the 95% standard line; Fig. 3.8), are clustered together in both the phenotypic and genotypic trees. 49 Oyster Water Sediment Figure 3.7. Vibrio spp. ERIC-PCR fingerprinting. Gel image showing the results of the ERIC-PCR on strains isolated in this study and additional strains for comparison. Strain designations are those listed in Table 3.2. Vibrio parahaemolyticus strains El-80 and E5-19 are included for reference since they were the strains used to isolate the viruses which generated the susceptibility patterns in Figure 3.6. Va: V. alginolyticus; Vp: V. parahaemolyticus; Vv: V. vulnificus. L: 100 bp ladders (100-1,500 bp + 2,000 bp top band). o 94.5% 0.1 Vp J00-6a Vp S00-1 Vp J00-5a Vp S00-2 Vp S00-5 Vp J00-9a Vp S00-3 Vp V497-B1 Vp J00-7a Vp V478-1 Vp C00-9a Vp V408-10 Vp V424-A3 Vp V416-7 Vp V465-A1 Vp V439-A1 Vp S00-8 Vp ATCC Vp E5-19 Vp E1-80 Va C00-6a Va S99-45 Va V478-16 Vp S01-31 Vp S01-20 Va ATCC Vv S99-48 Vp V p O V p R Va Figure 3.8. Vibrio spp. genotypic diversity. UPGMA tree of the Vibrio spp. genetic fingerprint similarities resulting from the gel image in Fig. 3.7. A Vibrio parahaemolyticus clade (Vp) and a V. alginolyticus clade (Va) are indicated, as well as two sub-groups consisting of oyster strains (VpO) and reference strains (VpR) within the Vp clade. Vibrio parahaemolyticus strains are identified as follows: black shaded (with white text) are from sediments, grey shaded are from oysters, and within boxes are from the water column. Strain designations are those listed in Table 3.2. Va: V. alginolyticus; Vp: V. parahaemolyticus; Vv: V. vulnificus. The scale bar represents a dissimilarity of 10%. The vertical line is the percent similarity (94.5%) of the identical standards (four 100 bp ladders) ran during the analysis - therefore, any groupings to the right of the line should be considered identical or very nearly so. 51 Comparisons can only be made to one study directly, and to two others indirectly (inference from presented data), as there are few researchers that have looked specifically at the relationship between phage-typing and genotypic diversity in the marine environment. The only direct study involved correlating Vibrio spp. phage susceptibilities with DNA-DNA hybridizations (Baross et al., 1978a). The two indirect studies examined phage sensitivity patterns as compared to host phylogenies of Pseudoalteromonas spp. (Wichels et al, 1998) and cyanobacteria (Sullivan et al, 2003). The overall interpretations generally match the data from this study - some larger phage-typing and genetic groupings of strains were evident, but all three studies found a general mismatch between the phage-typing and genetic diversity patterns. A final source for comparison, although only from the genetic stand-point, is the myriad of studies looking at the genotyping of V. parahaemolyticus strains (e.g. Wong et al, 1996; Marshall et al, 1999; Wong et al, 1999; Wong and Lin, 2001). However, these studies frequently look at clinical strains, at times compared to "environmental" strains, and have not examined the diversity of strains from differing environmental sources as in this report. Therefore, the novel patterns of diversity observed in this study must be discussed in the context of the distribution of Vibrio spp. in the environments investigated. 3.4.3 Distribution and Diversity Relationships The ubiquitous distribution of Vibrio spp. in the environments that were sampled suggests that they could play a significant role as hosts for viral infection within the bacterioplankton. In a study of Atlantic-Ocean bacteria, Moebus and Natkemper (1983) showed that 362 of 366 isolated phage-sensitive strains belonged to the Vibrionaceae family, also suggesting their contribution to virioplankton production. The wide range of environments where Vibrio spp. are found reflects the wide range of conditions that support their growth. The range of salinities, for example, in which Vibrio spp. occur ranges from near freshwater to -48 psu (Baumann and Shubert, 1984). Species such as V. cholera and V. parahaemolyticus thrive in estuarine environments (Grimes, 1991; Fukushima and Seki, 2004), consistent with the fairly high abundance of Vibrio spp. in the Fraser River Plume. The diverse array of Vibrio spp. that are pathogenic and/or commensal inhabitants of fish and shellfish (Muroga, 2001; Thompson et al, 2004a) may explain the slightly higher abundances of Vibrio spp. in areas of active aquaculture. Whereas there was no major trend between the phage-typing patterns of the Vibrio spp. strains and their geographic origins, there was separation based on the environments investigated. Vibrio spp. were much more abundant in the sediments, especially at the water-sediment 52 interface (up to 2.8 x 104 CFU cc"1), normally the most biologically productive zone of marine sediments (Eardly et al, 2001). As well, abundances of Vibrio spp. in coastal BC oysters (up to 7.5 x 104 CFU cc"1; Chapter II) were comparable to, if not higher than have been found in sediments. Abundances in the water column, by contrast, were much lower (only up to 346 mL" '). These trends were reflected in the susceptibility of isolates to viral infection. Sediments and oysters, where the highest abundances of Vibrio spp. occur, generally produced strains of V. parahaemolyticus that were permissive to viral infection; while the opposite was true for the water column. In oysters, the viral populations are abundant and dynamic, indicating the presence of susceptible host populations that support phage production (Chapter II). The situation in the sediment is less clear; vibriophage have not been detected in these sediments (Chapter V), however Vibrio spp. are abundant. If the bacteria are rapidly turning over, this should encourage viral infection if vibriophage are present. In the sediments, high abundances of Vibrio spp. and viruses (Maranger and Bird, 1996; Drake et al, 1998; Danovaro and Serresi, 2000) suggest conditions that would favour the presence of vibriophage and resistant cells, while low water-column abundances should favour susceptible cells, the opposite of what was observed. In the water column, low contact rates between Vibrio spp. and vibriophages (Paul et al, 1997; Chapter V) should not select for co-occurring resistance (Waterbury and Valois, 1993; Suttle and Chan, 1994), and should allow for the persistence of permissive bacteria. A possible explanation is that low abundances can lead to selection for lysogeny (reviewed in Wommack and Colwell, 2000), which would possibly make the water-column strains resistant to further infection by related VpVs (i.e.: super-infection prevention; Stewart and Levin, 1984). To test this hypothesis, the water-column strains (all but S01-31) were treated with mitomycin C to induce temperate phage. Four of the six strains tested did produce phage particles, however, the most permissive sediment and oyster strains also proved to be lysogens by the same method. This indicated that there was no correlation between resistance to phage and lysogeny. Whatever the cause of the resistance, the result is a group of water-column V. parahaemolyticus strains that are phenotypically (by phage-typing) more closely related to other members of the genus than to their own species. The genotypic diversity of the strains was less tied to the environment than were the susceptibility patterns. Although the fingerprinting reproduced the known divisions among Vibrio spp., there was no clear genetic separation between V. parahaemolyticus strains isolated from the sediment and water column, even though their susceptibility to phage infection is distinct. Strains from oysters, however, did cluster similarly in terms of phenotype and 53 genotype. This implies genetic homogeneity between V. parahaemolyticus in the sediment and their overlying waters, whereas genetic strains within the oysters are somewhat distinct. The typical isolation of primarily tdh~ (non-pathogenic) strains of V. parahaemolyticus from the environment, whereas tdh+ (pathogenic) strains primarily occur in shellfish (Shirai et al, 1990), provides further evidence of genetically distinct reservoirs. Therefore, the observation of different phage susceptibility patterns in genetically similar strains of V. parahaemolyticus from the water and sediment may result from phenotypic selection for different virus receptors. Alternatively, the genetic changes that affect virus receptors may be too small to be detected by genetic fingerprinting. For example, only very small changes in the LPS layer greatly affect susceptibility of E. coli to infection by T4 (Henning and Hashemolhosseini, 1994). 3.4.4 Conclusions The widespread presence of Vibrio spp. in sediments, oysters (Chapter II) and the water column show that they could support vibriophage populations in a variety of environments. Not surprisingly, therefore, geographic distribution was a poor determinant of diversity. The environment, however, had a much stronger effect on susceptibility to viral infection and, to a lesser extent, genetic diversity. This is the first known investigation of the distribution of V. parahaemolyticus genotypes and phenotypes (phage-types) in multiple environments. The results show that there is segregation of bacterial strains, under differing selection pressures, which will have an impact on in situ vibriophage production for which they act as hosts. The findings have general implications on bacterial-viral interactions in the pelagic environment. Although Vibrio spp. and their viruses (Chapter V) are endemic to the water column (as are many other marine hosts and viruses), it appears that virus production may be more prevalent in other environments, such as oysters (Chapter II) and possibly in sediments, given host abundances and susceptibility patterns. However, water-column viruses do persist in the 2 1 presence of low abundances of Vibrio spp. (maximum 10 mL" ) implying some production occurs in areas where the other sources are less prevalent. Contact rates between host and virus would be extremely low at these abundances, and a minimum of 103-104 host cells mL"1 is generally accepted as the experimental and observed threshold for successful lytic virus propagation (reviewed in Wommack and Colwell, 2000). A small amount of water-column production, along with larger inputs from other environments, may be a common way to maintain persistent virus populations of the many host species that are in low abundances (<10 mL"1) in the pelagic environment. 54 Chapter IV : The Use of Degenerate-Primed Random Amplification of Polymorphic DNA (DP-RAPD) for Strain-Typing and Inferring the Genetic Similarity Among Closely Related Viruses A version of this chapter had been published. Comeau, A.M., Short, S.M., and Suttle, CA. (2004) The use of Degenerate-Primed Random Amplification of Polymorphic DNA (DP-RAPD) for strain-typing and inferring the genetic similarity among closely related viruses. J Virol Meth 118: 95-100. 55 4.1 Summary Often it is necessary to distinguish among strains of closely related viruses, as well as infer the genetic relatedness within large groups of viruses. Current methods for strain typing viruses are time-consuming, require significant quantities of extracted DNA, and/or may require a priori genetic information. In this study, random amplification of polymorphic DNA (RAPD) was modified using a degenerate primer to produce unique and reproducible banding patterns from viral genomes. In the degenerate-primed RAPD analysis (DP-RAPD), a selection of algal virus and bacteriophage strains were profiled that encompassed an array of genome sizes and virus families. Closely related viruses (e.g. strains infecting Micromonas pusilla) generated similar, yet unique DP-RAPD patterns that could be readily distinguished from viruses within the same family (Phycodnaviridae) infecting a Chlorella-hke alga. As well, marine vibriophage from the families Myoviridae, Siphoviridae, and Podoviridae showed high diversity and were distinct from coliphage and cyanophage. Contamination of host DNA, even at levels above those that would normally be encountered, did not interfere with the viral patterns. These findings describe a rapid, PCR-based tool for strain-typing viral isolates that allows inferences to be made on genetic relatedness within groups of closely related viruses. 56 4.2 Introduction The field of marine viral ecology is accelerating rapidly as new applications of techniques yield efficient and novel approaches to research. Examples of these are the use of epifluorescence dyes and flow cytometry in the quantification of viruses (Hennes and Suttle, 1995; Noble and Fuhrman, 1998; Marie et al, 1999); and the use of denaturing gradient gel electrophoresis (DGGE) and pulse-field gel electrophoresis (PFGE) in the examination of viral diversity (Wommack et al, 1999a; Steward et al, 2000; Short and Suttle, 2002). Research in this field frequently results in the isolation of many viruses into culture, which can be close or distantly related to each other. Often, further information is desired on the degree of relatedness among these strains in order to assess geographic or temporal patterns of diversity or to make decisions on which isolates require further study (e.g. for pathogenesis studies, sequencing, etc.). The currently available methods for examining strain variation in viruses, such as restriction digests of genomic DNA and DNA/DNA hybridizations (Kellogg et al, 1995; Wommack et al, 1999b), are time-consuming and require significant quantities of extracted DNA. Other methods, such as DGGE and RFLP of gene products, require a priori genetic information on the strains in question. The lack of a rapid method for genetically fingerprinting uncharacterized viral isolates, led to the examination of the potential of modifying random amplification of polymorphic DNA (RAPD) for use on viral genomes. RAPDs are typically used for strain typing bacterial isolates (Wong et al, 1999) or for comparing whole bacterial communities (Franklin et al, 1999). However, the smaller genomes of viruses (1-10% that of bacteria) prevents using the standard RAPD technique, as the typical sequence-specific 10-mer random primers result in too few bands. In this study, an optimized degenerate-primer RAPD assay (DP-RAPD) was used to profile algal virus and bacteriophage isolates, encompassing a range of genome sizes, virus families, and hosts (Table 4.1). The utility of the assay for producing unique and reproducible fingerprints from viral genomes is demonstrated. 57 Table 4.1. Selected characteristics of the virus strains used in this study. Virus Strain" Host Genome Size (kb) Virus Family Sourceb References Algal viruses MpV GM1 Micromonas pusilla 200c Phycodnaviridae Gulf of Mexico, TX Cottrell and Suttle, 1991 MpV SP1 Micromonas pusilla 200c Phycodnaviridae Scripps pier, CA Cottrell and Suttle, 1991 MpV PB5 Micromonas pusilla 200c Phycodnaviridae Peconic Bay, NY Cottrell and Suttle, 1991 MpV PB6 Micromonas pusilla 200° Phycodnaviridae Peconic Bay, NY Cottrell and Suttle, 1991 MpV PL1 Micromonas pusilla 200c Phycodnaviridae Aransas Pass, TX Cottrell and Suttle, 1991 ChiV CVA-1 Chlorella sp. 330 Phycodnaviridae J. Van Etten Van Etten and Meints, 1999 ChlV PBCV-I Chlorella sp. >300 Phycodnaviridae J. Van Etten Van Etten and Meints, 1999 Bacteriophage VpV 262 Vibrio parahaemolyticus 45.9 Podoviridae Strait of Georgia, BC Hardies etal., 2003; Chapter V VpV 395 Vibrio parahaemolyticus 107 Siphoviridae Strait of Georgia, BC Chapter V VpV 396 Vibrio parahaemolyticus 79 Siphoviridae Strait of Georgia, BC Chapter V VpV 404 Vibrio parahaemolyticus 65 Siphoviridae Strait of Georgia, BC Chapter V S-PWM2 Synechococcussp. 200 Myoviridae Gulf of Mexico, TX Suttle and Chan, 1993 S-PWM3 Synechococcussp. 200 Myoviridae Gulf of Mexico, TX Hambly. 2002; Suttle and Chan, 1993 S-PWM4 Synechococcussp. 200 Myoviridae ' Gulf of Mexico, TX Suttle and Chan, 1993 X ' Escherichia coli 48.5 Siphoviridae Commercial supplier -T4 Escherichia coli 169 Myoviridae E. Kutter -T7 Escherichia coli 39.9 Podoviridae H. Ackermann -a MpV: Micromonas pusilla Virus; ChlV: Chlorella Virus; VpV: Vibrio parahaemolyticus Virus. b Viruses with geographic sources were isolated in the Suttle lab, otherwise they were provided as listed. c The size of MpV strains reported in Cottrell and Suttle (1991) is incorrect and was determined for MpV-SPl to be ca. 200 kb by PFGE (unpublished). The genome sizes of the other MpV strains has not been determined but is also assumed to be ca. 200 kb based on RFLP analysis. 4.3 Materials and Methods 4.3.1 Viral DNA Bacteriophage lysates were prepared by the plate lysate method (Chapter II; Sambrook et al, 1989) and were 0.22 pm-filtered to remove any remaining host cells and cellular debris. For the algal viruses (except CVA-1), aliquots (100 (iL) of filtered lysates (provided by S. M. Short) were subjected to a hot-cold treatment to release the viral DNA (Chen et al, 1996) - 3 cycles of 95 °C for 2 min and 4 °C for 2 min; 3 pL of the extract was used in each reaction. At least a 10-fold dilution of the extracts was generally needed to obtain clear PCR products because of the interference of salts present in the marine media used to amplify the viruses. For the Chlorella Virus (CVA-1), 30 ng of DNA (provided by J. Van Etten) was used in the assay instead of hot-cold treated lysate. Bacteriophage DNA was isolated using a phenol:chloroform extraction and 30 ng of this template were also used in reactions. A mixed template reaction, containing four viruses in equal proportion, also contained a final amount of 30 ng. Bacteriophage X DNA was obtained commercially (Promega). 4.3.2 Host DNA In order to determine the extent of interference from host DNA contamination, varying amounts (2-200 ng) of Vibrio parahaemolyticus DNA (strain 94Z944 El-80 - BCCDC; phenol:chloroform extraction) were included in reactions with V. parahaemolyticus Virus 396 (VpV 396; 20 ng). The copy number of host DNA was calculated using the base-pair weight of DNA and the length of the V. parahaemolyticus genome (5.2 Mbp; Makino et al, 2003). 4.3.3 DP-RAPD PCR The 50 p:L DP-RAPD reactions consisted of the following reagents (Invitrogen Life Technologies): Taq DNA polymerase buffer, 4.5 mM MgCl2, 0.6 mM of each deoxyribonucleoside triphosphate, 2 pM of the R10D primer, and 1.5 U of PLATINUM Taq DNA polymerase. Negative controls contained all reagents except template DNA. The R10D (RAPD 10-mer degenerate) primer sequence was 5'-GTCASSWSSW-3' where S and W represent G/C and A/T, respectively. This sequence was chosen in order to begin with an unbiased specific run of bases (i.e.: one each = GTCA), followed by 6 bases of 2-fold degeneracy (64-fold total) in order to be in the range of the 10- to 100-fold genome size difference between bacteria and viruses. The double repeat of the degenerate section was 59 selected in order to have an increased G/C content for stability (i.e.: more S versus W), but to prevent biased amplification of runs of any one base in template sequences (i.e.: avoid three or more concurrent similar bases). PCR was carried out with the following cycle parameters: initial denaturation at 95°C for 1.5 min; followed by 40 cycles of denaturation at 95°C for 45 s, annealing at 40°C for 3 min, extension at 72°C for 1 min; and a final extension at 72°C for 10 min. PCR products (10 p:L each) and 100 bp ladders (Invitrogen Life Technologies) were electrophoresed in 1.5% agarose (Invitrogen Life Technologies) in 0.5X TBE buffer (45 mM Tris-borate, 1 mM EDTA [pH 8.0]) at 90 V for 3 h. Gels were stained with ethidium bromide and captured with a Chemilmager 5500 gel documentation system (Alpha Innotech Corporation). 4.3.4 Fingerprint Analysis Digital gel images were cropped and enhanced (brightness/contrast) using Adobe Photoshop 5.0 LE. Fingerprints were compared quantitatively using GelCompar II (Applied Maths). Bending in the gels, which occasionally leads to incorrect visual assessment of fingerprint similarities, was corrected for in the software before analysis. To quantify relationships among strains, UPGMA trees were constructed using a Pearson correlation method that compares the densitometry profiles of each lane to one another. For a more in-depth explanation of the GelCompar analysis, consult Ferrari and Hollibaugh (1999). Similarity values were normalized for the variability within gels by adding to them the differences between two identical standards (5.2-7.1%; 100 bp ladders) run on each gel. 60 4.4 Results and Discussion 4.4.1 Optimization of the DP-RAPD PCR The use of a short, degenerate oligonucleotide in the DP-RAPD assay required optimization of PCR conditions. Initial trials with 6- and 8-mer random primers resulted in no product formation, even at very low-stringency conditions, presumably due to the disassociation of the primers from the template DNA once the reaction temperature was increased to 72°C for the extension step. The R10D primer however, resulted in reproducible and distinct banding patterns (Fig. 4.1 and 4.2) using the final conditions described. The MgCl2 concentration was increased relative to standard PCR and the annealing temperature of 40°C was held for 3 min in each cycle to stabilize the primer-template complexes. The primer was also designed with a G/C content of 60% to add to complex stability. Due to the increased number of target sites in random PCR, the concentrations of primer, nucleosides, and polymerase enzyme were also increased. Even greater concentrations (and lower stringency) were tested, including a gradient of template concentrations. However, fingerprints began to smear and/or were inconsistent above or below the final stringency conditions listed above (data not shown). For instance, template amounts above 30 ng resulted in a loss of band resolution due to bands becoming overloaded, whereas many fainter bands disappeared at amounts below 30 ng. It was also found that DNA template preparation from algal viruses only required hot-cold lysis, whereas phenol-chloroform extraction was required for bacteriophage. A comparison of the two methods can be seen in Figure 4.1; the CVA-1 fingerprint (lane 6) was generated from isolated DNA, whereas the PBCV-1 fingerprint (lane 7) was from a hot-cold lysis. 4.4.2 DP-RAPD Virus Fingerprints The algal virus DP-RAPD fingerprints (Fig. 4.1 A) showed that the Micromonas pusilla Virus (MpV) strains were all unique, yet shared closely related banding patterns. It is also quite clear that they are distinct from the Chlorella viruses (ChlVs) PBCV-1 and CVA-1, which also demonstrated very similar fingerprints. These results are consistent with the relationship between MpV and ChlV strains based on DNA polymerase (pol) gene phylogeny and whole genomic hybridization data (Cottrell and Suttle, 1995; Chen and Suttle, 1996). The fingerprints indicated that the minimum similarity among MpV strains averaged 58 % (Fig. 4. IB) compared to 78% for DNA pol and 20% for hybridization. The similarity between the MpVs and ChlVs based on fingerprint analysis was 26%, compared to 56% based on sequence analysis of pol gene 61 fragments and undetectable based on hybridization. Based on these data, the sensitivity of the DP-RAPD assay to resolve genetic differences lies between sequence analysis of pol genes and DNA hybridization. When the two ChlVs are compared, their DP-RAPD fingerprints were remarkably similar (98%), whereas their degree of relatedness from DNA pol data is 77% (hybridization data are not available). This disparity may be the result of sequence analysis reflecting single gene evolution, whereas the DP-RAPD assay represents a more "genomic" measure. Further experimentation or genome sequencing may demonstrate whether PBCV-1 and CVA-1 are more closely related than is implied by their DNA pol sequences. Another possible explanation of the higher similarity between the ChlVs may be that as the number of bands in the DP-RAPD profiles increases there is a greater probability that co-migrating bands will occur by random chance, thereby increasing the similarity values. This would tend to occur more frequently in larger genomes, as is the case with the ChlVs (>300 kb). It is suggested that, when analyzing very large (or, conversely, very small) viral genomes, the conditions reported in this study should be used as the starting point to optimize banding patterns. The bacteriophage fingerprints (Fig. 4.2A) demonstrated a much greater degree of diversity than was observed for the algal viruses. This was expected since the phage comprised three virus families, instead of one, and a large range of genome sizes (39.9-200 kb; Table 4.1). The fingerprints were generally composed of fewer bands (mean = 10.6, range = 7-16) than the larger algal virus fingerprints (mean = 14.1, range = 11-19), but there was only a moderate linear relationship between genome size and the number of bands (r2 = 0.57; data not shown). The similarity tree (Fig. 4.2B) showed that most of the viruses were only distantly related. The only exceptions to this trend were the cyanomyoviruses (S-PWM2/3/4) that are known to be related (similar morphologies and genome sizes; Suttle and Chan, 1993) and which clustered at a similarity level of 90%. The only other myovirus among the phage examined was T4 and it was the closest fingerprint to the cyanophage. The siphoviruses clustered around the myoviruses to form a large group whose fingerprints were only distantly related to the two podoviruses examined in this study (T7 and VpV 262). Recent sequencing information (Hardies et al, 2003) agrees with the large degree of divergence (< 30% DP-RAPD similarity) between the T7 and VpV 262 fingerprints. Although the fingerprints reflected general virus groupings, the goal was not to analyze the genetic distance among distantly related viruses, but to demonstrate that the DP-RAPD assay can be used on a wide variety of virus types. The most appropriate use of the assay is to make comparisons within groups of related viruses (i.e.: strain typing). 62 MpVs B 20 40 60 80 i i i i i i i i i i i i i i 100 — SP1 PB6 PB5 PL1 GM1 r CVA-1 PBCV-1 < o < Figure 4.1. Analysis of algal virus genomes. (A) Agarose gel of DP-RAPD fingerprints for strains of Micromonas pusilla Viruses (MpVs - lanes 1-5) and Chlorella sp. Viruses (ChlVs -lanes 6-7). Strains that were examined are as follows: Lane 1: GM1, lane 2: SP1, lane 3: PB5, lane 4: PB6, lane 5: PL1, lane 6: CVA-1, lane 7: PBCV-1, L: 100 bp ladder with selected marker sizes to the left of the gel image. (B) UPGMA tree of the fingerprints in panel A with % similarity scale bar. 63 Vibriocb Cyano(|) B 30 40 60 • i • 80 • i i • 100 L _ J - VpV 395 - VpV 404 S-PWM2 S-PWM3 S-PWM4 T4 VpV 396 X T7 VpV 262 o CO •o' 3" o Tl O a o Figure 4.2. Analysis of bacteriophage genomes. (A) Agarose gel of DP-RAPD fingerprints of vibriophages (Vibriocj) - lanes 1-4), cyanophages (Cyano(() - lanes 5-7), and coliphages (Coli(b -lanes 8-10). Strains that were examined are as follows: Lane 1: VpV 262, lane 2: VpV 395, lane 3: VpV 396, lane 4: VpV 404, lane 5: S-PWM2, lane 6: S-PWM3, lane 7: S-PWM4, lane 8: X, lane 9: T4, lane 10: T7, L: 100 bp ladder (sizes as per Fig. 4.1 A). (B) UPGMA tree of the fingerprints in panel A with % similarity scale bar. Myo / Sipho = myoviruses and siphoviruses, Podo = podoviruses. 64 4.4.3 Template Mixtures in the DP-RAPD Given that the DP-RAPD assay uses random primers and operates under low stringency conditions, there was interest in determining how the assay would respond to template mixtures. The two scenarios tested were: 1) an equal mixture of virus templates; and 2) contamination of one viral template with varying levels of host bacterial DNA. The results of both scenarios are presented in Figure 4.3. In the first scenario, it was evident from the fingerprints that the viruses (lanes 1-4) did not contribute equally to the profile based on the mixture of phage (lane 5). There was a tendency for some strong bands to be retained and some weaker bands to be lost. This did not happen evenly among templates, nor was there a relationship between genome size and the contribution of each virus. The similarity values between the viruses and the mixture fingerprint (which measure the relative contributions of each virus to the mixture) ranged from 32-66%. VpV 404 (lane 4) contributed the most to the mixed fingerprint, followed by VpV 396 (lane 3), VpV 395 (lane 2), and VpV 262 (lane 1). Presumably, PCR biases at certain loci, which already result in differing band intensities within a template, led to the unequal contributions of templates in the mixture. Consequently, PCR biases and the severe dilution of individual templates in complex communities would prohibit using the assay to compare similarities of natural viral assemblages. However, the assay could complement a method such as PFGE of whole virus community DNA where it would enable discrimination of differing virus types that migrate to the same location (i.e.: similar genome sizes). The second scenario was designed to test how sensitive the DP-RAPD assay would be to contamination of viral template by host DNA. Vibrio parahaemolyticus DNA produced a faint ladder-like banding pattern at 20 and 200 ng of template per reaction (Fig. 4.3). These amounts represent copy numbers of 4 x 106 and 4 x 107; and are the equivalent of ~109 and ~1010 cells mL"1 of contamination in a viral lysate (assuming 100% extraction). The host DNA was then mixed with 20 ng of virus DNA (VpV 396), equivalent to 2.5 x 108 copies or -1011 viruses mL"1. There were only minor effects in the 650 bp range on the virus fingerprint at the highest level of contamination (200 ng). There was no effect at the two lower levels of contamination, presumably because the copy number favours the viral template to a much greater degree. As these levels of DNA contamination are well above amounts that would likely occur in prepared viral lysates, the DP-RAPD assay should be fairly robust in the presence of host DNA contamination. 65 Virus DNA (ng) 20 20 20 Host DNA (ng) 1 2 3 4 5 L 200 20 2 200 20 2 Virus DNA (ng) 20 20 20 Figure 4.3. Influence of template mixtures on fingerprints. DP-RAPD fingerprints of four viral templates (lanes 1-4; VpVs 262, 395, 396, and 404) and an equal mixture of all four templates (lane 5). L: 100 bp ladder (sizes as per Fig. 4.1 A). The final six lanes are host and host + virus fingerprints. Host DNA is Vibrio parahaemolyticus and virus DNA is VpV 396. DNA amounts are listed in nanograms (ng). 66 In summary, the DP-RAPD assay presented in this study resulted in unique and reproducible fingerprints generated from an array of algal viruses and bacteriophages. The assay was useful in discriminating between closely related strains and generally reproduced the patterns of relatedness within virus families. Results from mixtures of viral templates indicate that the assay would probably be of limited use in profiling complex natural viral communities, unless it complemented existing methods such as PFGE. However, the assay was fairly robust in the presence of host template contamination and should provide an excellent method for strain-typing and assessing the diversity of groups of novel viruses. The assay has been employed in generating random sequence from novel viruses and for profiling large groups of novel vibriophage (Chapter V). 67 Chapter V : Distribution and Diversity of Vibriophages in a Coastal Environment 68 5.1 Summary Although many marine viruses have been characterized, little is known about the distribution and diversity of single-species viruses in nature. The purpose of this study was to isolate and characterize Vibrio parahaemolyticus viruses (VpVs) from the Strait of Georgia, and to examine their spatial variation in host-range and genetic diversity. Nearly all enrichment cultures (29/31) generated isolates, implying that VpVs were nearly ubiquitous in the virioplankton, yet at very low abundances (<1 L" 1). Viruses were not detected in sediments (n = 99), albeit at a poorer sensitivity of detection. Fourteen of the 16 viruses characterized had long non-contractile tails (siphoviruses), with genome sizes ranging from -45-106 kb, and half were capable of infecting other Vibrio species. The VpVs infected bacteria from oysters and sediment fairly well (55 and 46% of the total 112 and 80 host-virus combinations, respectively), but were unable to infect many of the bacteria from the water column (<13% of 112 combinations). When compared to VpVs from oysters, it was clear that the major determinant of host-range and genetic diversity was not geography, but the source environment from which the VpVs originated. Viruses from some of the farthest separated locations (-120 km) were closely related, while the diversity within some locations was very high. Therefore, the VpV population within the Strait of Georgia is a highly diverse, yet homogeneous mixture of phenotypes and genotypes. 69 5.2 Introduction The virioplankton, numbering an estimated >10 viruses in the biosphere (Hendrix et al, 1999; Briissow and Hendrix, 2002), exert significant influence on marine phytoplankton and bacteria (reviewed in Wommack and Colwell, 2000). These effects include the reduction of productivity through viral-mediated mortality (Hewson et al, 2001; Wilhelm et al, 2002), the short-circuiting (virus shunt) of organic matter back into the dissolved pool through lysis products (Wilhelm and Suttle, 1999), mediating horizontal genetic exchange (Chibani-Chennoufi et al, 2004) and lysogenic conversion (Waldor and Mekalanos, 1996), and influencing the richness and evenness (i.e.: overall diversity) of host populations (Wommack et al, 1999b; Thingstad, 2000). Although the latter category of effects may be responsible for a significant portion of marine microbial structure, little information is available on the diversity of single host-virus systems in nature. I am interested in the spatial and temporal patterns of abundance, phenotype, and genotype of single-species host-virus systems. Although advances have been made in the study of single-species populations in natural samples, these have normally been restricted to the distribution, not the diversity (or correlations between the two), of hosts and viruses. A few noticeable exceptions were investigations on the genetic distribution of vibriophage (Kellogg et al, 1995), the co-variation of phycoviruses with eukaryotes (Short and Suttle, 2003), and the clonal composition of a phytoplankton host and its viruses during bloom conditions (Tarutani et al, 2000). There is a need for more of these types of investigations because it is still not fully known at which periods, and to what extent, viruses affect (or respond to) the structure (richness and evenness) of their host communities. A number of Vibrio parahaemolyticus viruses (VpVs) have been isolated and characterized from the coastal waters of Japan, Laos, Hawaii, Florida, and the Pacific and Atlantic coasts of North America (Nakanishi et al, 1966; Sklarow et al, 1973; Baross et al, 1974; Kellogg et al, 1995; Nakasone et al, 1999; Hardies et al, 2003; Seguritan et al, 2003). All isolates, except a few filamentous phages (Inoviridae family), belong to the three families of tailed phages, Myoviridae, Siphoviridae, and Podoviridae (Ackermann et al, 1984). The viruses are usually species-specific and sometimes strain-specific, although a few demonstrate broader host ranges against Vibrio and the related genus Photobacterium (Nakanishi et al, 1966; Baross et al, 191 A; Matsuzaki et al, 1992; Matsuzaki et al, 1998). Mitomycin C and ultraviolet light have been used as inducing agents in isolations of temperate VpVs (Ohnishi and Nozu, 1986; Koga and 70 Kawata, 1991) and one study found that -10% of V. parahaemolyticus isolates from various sources harboured lysogens (Muramatsu and Matsumoto, 1991). Although many VpVs have been characterized in terms of morphology, physical properties, and host range, little is known about their abundance and distribution in nature. A few studies have found up to 2.0 PFU mL"1 of VpVs in coastal waters although titers near 0.05-0.4 PFU mL"1 are more typical (Kellogg et al., 1995; Paul et al., 1997). In contrast, titers can reach 105-106 PFU g"1 (Baross et al., 1974; Baross et al., 1978b; Chapter II) in mollusc tissues. Abundances of VpVs in sediments are relatively unknown, with only one report of 7 PFU cc"1 in a Florida reef (Paul et al, 1993) and only one isolate from the Atlantic coast (Sklarow et al, 1973). Given the lack of basic distribution data, there is also little information on the distribution of specific phenotypes and genotypes in the marine environment. However, given that VpVs and V. parahaemolyticus are relatively well studied (compared to all other heterotrophic phage-host systems) and widely distributed (Koga et al, 1982; Kellogg et al, 1995; Thompson et al, 2004a), makes VpVs and their hosts an excellent model for examining phage diversity in nature. The purpose of this study was to isolate and characterize VpVs from the Strait of Georgia, with the final goal of examining host-range and genetic diversity in a spatial context. In the larger context of marine viruses, this will allow us to determine whether virus diversity patterns match host diversity patterns (Chapter III), and to see what implications these patterns have on virus production and host structure. 71 5.3 Materials and Methods 5.3.1 Sample Collection and Processing Water and sediment samples were collected from the Strait of Georgia, British Columbia and adjacent inlets (Fig. 5.1) between 1996 and 2003 (Table 5.1). Twenty to 200 L of water were collected using Go-Flo bottles mounted onto a rosette, or by surface sampling using a bucket (Jericho Pier only). The viral fraction was concentrated as outlined in Suttle et al. (1991). Briefly, seawater samples were pre-filtered through duplicate, 142 mm glass-fibre (1.2 pm GC50, MFS) and polyvinylidene fluoride filters (0.45 pm HVLP, Millipore) in series. The viral size fraction of the filtrate was then concentrated -100 (20 L samples) to -500 times (200 L samples) using a 30 kDa cut-off, spiral-wound ultrafiltration cartridge (Millipore). The ultrafiltrate (bacteria/virus-free) was retained as a diluent and base for media. Sediment-core samples were collected using a triple-barreled gravity corer (Rigosha and Co.) and pore waters extracted as described previously (Lawrence et al., 2002). Briefly, sediment-water interfaces were sampled using serological pipettes and deeper samples were sliced from the extruded cores. Twenty cm of each sample were mixed with an equal volume of PBS (Sambrook et al., 1989) in a 50 mL polypropylene tube and agitated on a tabletop shaker for 30 min at 150 rpm. Samples were then centrifuged at 4000 x g for 5 min at 4°C to separate coarse sediment materials from pore waters. The volumes of extruded pore waters and remaining sediments were recorded, and the supernatants were filtered through glass-fibre (1.2 pm GF/C, Whatman) and polyvinylidene fluoride filters (0.45 pm HVLP, Millipore). All processed liquid samples were then stored at 4°C in the dark until viral screening. Table 5.1. Detection of VpVs in samples from the Strait of Georgia, BC. Sample Type Isolation Method3 Sample Years Number of samples Total Positive Detection Limit" Water column Plaque 1996-2001 117 1 0.02-10 mL"1 Water column Enrichment 1996/1999/2000 31 29 0.05-0.9 L _ l Sediment pore water Plaque 1999/2000 99 0 7.5-123 cc-1 Oyster homogenate0 Plaque 2000/2001/2003 22 22 100 g 1 a See Experimental procedures for further details on isolation methods. b Assuming one infective virus in the assayed aliquot resulting in a single plaque or lysis of a liquid culture. 0 See Chapter II for further details on isolations from oysters. 72 5.3.2 Growth Media and Conditions All bacteria and viruses were suspended in ultrafiltered (see above), sterile natural seawater diluted to 15 psu with reverse-osmosis water (Millipore). The growth medium was Marine Luria-Bertani broth (MLB; 0.5 g L"1 each of casamino acids, peptone, and yeast extract, 0.3% v/v glycerol, in 15 psu ultrafiltrate base). This broth was amended with 1% w/v agar for solid plates and 0.6% w/v agar for plaque-assay overlays. Overnight cultures and enrichment assays were grown at 30°C with agitation (150-200 rpm). Plaque assay and bacterial stock plates were grown at room temperature (~25°C). 5.3.3 Vibrio parahaemolyticus Virus (VpV) Isolation and Amplification Viruses were isolated using liquid enrichment cultures (water column only) and plaque assays (Suttle, 1993). In enrichment assays, 9 volumes of concentrated viral communities (9-45 mL) were added to 1 volume of exponentially growing host cells (1-5 mL) and the cultures were monitored for lysis for up to 10 days. Positive (and negative) enrichments were confirmed by a subsequent plaque assay using aliquots of the cleared (or still turbid) lysates. Nine of the 31 water-column enrichments were composites of equal volumes taken from 3 to 10 viral community samples. Six of the nine were composites from similar locations (HS, JPx2, MI, PS, SA/SI; Fig. 5.1) and the remaining three were composites of depth profiles from single locations (HS, MI, SA; Fig. 5.1). Water-sample plaque assays used 5-100 pL of viral concentrate in (or on) an agar-overlay of host cell lawns. Given the limited volume available, only 15-100 p i of pore water was used for each sediment sample screening (plaque assays only). The volumes screened by plaque assay varied because the samples were either mixed with host cells in the plaque assay lawns (100 pL volume) or spotted onto poured lawns containing host cells only (5-20 uL spots). Viruses were plaque purified for 3 rounds, then amplified by the plate-lysate method (Sambrook et al., 1989). Briefly, several agar-overlay plates were created with the appropriate dilution of phage to obtain a completely lysed host lawn. Fifteen mL of sterile seawater were added to each plate, the thin lawn was shredded using a sterile loop (to create more contact area), and the plates were allowed to elute at 4°C overnight. The agar/seawater mixture was then transferred to a sterile tube, vortexed, and centrifuged at 4100 x g for 60 s to sediment large debris. Finally, the resultant lysate supernatants were 0.22 pm-filtered to remove any remaining bacteria and maintained at 4°C in the dark. The two Vibrio parahaemolyticus strains used for screening, clinical isolate 94Z944 El-80 and environmental isolate 98S128 E5-73 19, were determined to be the most permissive hosts (of an examined subset, n = 22) available from the BC Centre for Disease Control, based on PFUs recovered from natural samples. Viruses first isolated from oysters (Chapter II) were further characterized (PFGE, electron microscopy, and genetic fingerprinting) in this work in order to provide comparison strains to the water-column viruses. 5.3.4 Pulse Field Gel Electrophoresis (PFGE) Genome sizes were determined by extracting the viruses in TEGED (10 mM Tris, 1 mM EGTA, 1 mM EDTA) in preparation for PFGE (Steward et al, 2000). Aliquots of concentrated lysate were incubated with IX TEGED at 65°C for 30 min, and then run on 1% agarose gels in 0.5X TBE (45 mM Tris-borate, 1 mM EDTA [pH 8.0]) using a CHEF Mapper XA (Bio-Rad) at 6.0 V/cm for 16 h, with an included angle of 120°, an initial switch time of 1 s and a final switch time of 6 s at a ramp rate of 0.357. Gels were stained with SYBR Gold (Molecular Probes) and captured with an Alphalmager 3400 gel documentation system (Alpha Innotech Corporation). Approximate genome sizes were determined using reference standards (Bio-Rad) and software analysis (AlphaEaseFC, Alpha Innotech Corporation). 5.3.5 Electron Microscopy Concentrated lysates were adsorbed onto formvar-coated grids (400 mesh; Ted Pella) for 0.5-2 h, wicked dry using filter paper, stained with 1% uranyl acetate for 30 s, and wicked dry for a final time. The grids were observed with a transmission electron microscope (Hitachi H7600) at an accelerating voltage of 80 kV. 5.3.6 Host Range Assays Host ranges were determined using strains of V. parahaemolyticus, as well as other strains of Vibrio spp., isolated from BC coastal waters and other areas (Table 5.2). Ten to twenty \ih of amplified virus stocks were spotted into agar-overlay lawns of bacteria and monitored for zones of clearing for 1-7 d. The results were scored as presence (zone of lysis = "1") or absence (no lysis = "0") data. Host ranges of viruses isolated from oysters (Chapter II) were included in the analysis. A distance matrix was then calculated using the Dice similarity coefficient (2nab / [na + nb]; Dice, 1945), which is basically the ratio of things in common, out of the total, between each pair-wise combination of viruses. Twice the number of shared elements (2nat,; in this case shared "0s" and "Is") between any two viruses is divided by the total number of elements in both 74 viruses (na + nt,; total number of "Os" and "Is"), resulting in a value between 0 and 1. This distance matrix was then used to create a UPGMA tree using the NEIGHBOR function in PHYLTP v3.5 (http://evolution.genetics.washington.edu/phylip.html). Five of the 32 tested host strains were removed from the analysis since they were negative for all viruses and therefore contained no resolving information. Table 5.2. Source of bacterial strains used to determine the host range of VpVs. All strains were isolated during this dissertation (Chapter III), except for ATCC strains and those with a " V " strain designation (see Chapter II). Species Strain(s) Source Vibrio parahaemolyticus V408-10to V497-B1 Ladysmith Harbour, BC Vibrio parahaemolyticus C00-9a Ladysmith Harbour, BC Vibrio parahaemolyticus JOO-5 to J()0-9a Jericho Pier, BC Vibrio parahaemolyticus SOO-1 to SO 1-31 Strait of Georgia, BC Vibrio alginolyticus ATCC 17749 ATCC" Vibrio alginolyticus C00-6a Ladysmith Harbour, BC Vibrio alginolyticus .100-Ida Jericho Pier, BC Vibrio alginolyticus l'\VII3a Gulf of Mexico, TX Vibrio alginolyticus S99-45 / S00-4 Strait of Georgia, BC Vibrio alginolyticus V478-16 to V497-C16 Ladysmith Harbour, BC Vibrio fluvialis S99-44 Strait of Georgia, BC Vibrio natriegens ATCC 14048 ATCC Vibrio vulnificus S99-48 Strait of Georgia, BC Vibrio vulnificus S01-38 Strait of Georgia, BC 5.3.7 Genetic Fingerprinting Genetic fingerprinting was carried out as described for phage in Chapter IV with only slight modifications. A selection of the viruses (8 of 29) required the addition of 5% (final concentration) dimethyl sulfoxide (DMSO, Sigma), which reduces secondary structure, to the PCR reactions in order to obtain strong, reproducible fingerprints. Finally, PCR products were electrophoresed at 110 V for 3 h and the gel image was captured with an Alphalmager 3400 gel documentation system (Alpha Innotech Corporation). The phages T4 (provided by E. Kutter), T7 (provided by H. Ackermann), and lambda (Promega) were included as standards for comparison. 75 5.4 Results and Discussion 5.4.1 Presence of VpVs in the Strait of Georgia Isolation of Vibrio parahaemolyticus viruses (VpVs) from inlets adjacent to the Strait of Georgia (Fig. 5.1) depended upon the environment examined and the method employed (Table 5.1). Plaque assays of 117 water samples yielded only one positive (detection limit: 0.02-10 mL" l), whereas 29 of 31 enrichment cultures (detection limit: 0.05-0.9 L"1) generated isolates. It is assumed that the much greater success of the enrichments is due to the -1000-fold greater sensitivity of the latter assay. These data imply that VpVs are common but typically at low abundances (<1 L"1) in the Strait of Georgia region. Similar water-column studies have found mean abundances between 10 and 400 PFU L"1 in coastal Florida and Hawaii (Kellogg et al., 1995; Paul et al., 1997), indicating that a low background abundance of VpVs may be typical of marine waters. This situation is similar for viruses infecting the photosynthetic flagellate Micromonas pusilla and cyanobacteria of the genus Synechococcus which are widespread throughout the world's oceans (Cottrell and Suttle, 1991; Suttle and Chan, 1993; Waterbury and Valois, 1993; Wilson et al, 1993; Suttle and Chan, 1994; Cottrell and Suttle, 1995; Sahlsten, 1998; Zingone et al, 1999; Ortmann et al, 2002). VpVs were undetectable in 99 sediment samples taken from inlets and bays adjacent to the Strait of Georgia, at depths from the water-sediment interface down to 40 cm (Table 5.1). Although the detection limits in sediments were relatively insensitive (8-123 cc"1), the high abundances of host cells (Chapter III) and of virus particles in marine sediments (Drake et al, 1998; Danovaro and Serresi, 2000), suggested they might be readily detectable. Moreover, viruses infecting eukaryotic phytoplankton (Lawrence et al, 2002) and cyanobacteria (Suttle, 2000) have been found down to 40 cm in sediment cores, some of which were collected at the same locations as the samples assayed for VpVs in this study. Yet the only reports of VpVs in sediments are a report of 7 PFU cc"1 in a sediment sample from Florida (Paul et al, 1993) and an isolate from the coast of North Carolina (Sklarow et al, 1973). The low, to undetectable, abundances in water and sediments is contrasted by the ease of isolation, and very high abundance, of VpVs in oysters (Baross et al, 1974; Baross et al, 1978b; Chapter II). For example, regardless of season, oysters in Ladysmith Harbour (LH, Fig. 5.1) contained -0.5-10 x 104 VpVs cc"1 (Chapter II). 76 50°N 49° N 125°W 124°W 123°W F i g u r e 5.1. G e o g r a p h i c d i s t r i b u t i o n o f V p V s . Locations positive for the presence of VpVs are indicated with closed circles (•). FRP: Fraser River Plume; HS: Howe Sound; JP: Jericho Pier; LH: Ladysmith Harbour; MI: Malaspina Inlet; NI: Narrows Inlet; PS: Pendrell Sound; SA: Salmon Inlet; SI: Sechelt Inlet. 77 5.4.2 VpV Morphologies and Genome Sizes Most VpVs were siphoviruses (long, non-contractile tails), including four strains with elongated heads (B2 morphotype) (Table 5.3, Fig. 5.2), although podoviruses were also common. To my knowledge, these are the first podoviruses (which include VpV 262 [Hardies et al, 2003]) that have been isolated infecting V. parahaemolyticus. Most described VpVs have been myoviruses (Muramatsu and Matsumoto, 1991; Matsuzaki et al, 1992; Kellogg et al, 1995; Matsuzaki et al, 1998; Matsuzaki et al, 2000), although a number of siphoviruses (Hidaka and Tokushige, 1978; Koga et al, 1982; Koga and Kawata, 1991) and a few filamentous inoviruses (Nakanishi et al, 1966; Taniguchi et al, 1984; Nakasone et al, 1999; Nasu et al, 2000; Chang et al, 2002) have been reported. Skewed distributions of morphotypes also occur with viruses infecting marine cyanobacteria and prochlorophytes, most of which are myoviruses (Suttle and Chan, 1993; Waterbury and Valois, 1993; Wilson et al, 1993; Sullivan et al, 2003). In general, vibriophages appear to be more evenly distributed among the three tailed-phage families (Ackermann et al, 1984; Pelon et al, 1995; DePaola et al, 1998). Therefore, it is unknown whether VpVs are atypical in their distribution, whether previous isolations have unwittingly selected for myoviruses, or whether this study selected against them. The dominance of one morphotype over another may be a moot point given the temporal and spatial variability in virioplankton communities (Bratbak et al, 1990; Hennes and Simon, 1995). For example, myoviruses dominated North Sea and North Atlantic communities (Frank and Moebus, 1987), while siphoviruses and podoviruses were the most abundant in Lake PluBsee (Demuth et al, 1993), and the northwest Pacific (Hara et al, 1991), respectively. A more recent analysis of two whole-community shotgun libraries found slightly more podovirus-like sequences in one sample and an equal amount to siphoviruses in the other (Breitbart et al, 2002). The distribution of morphotypes is of interest since it may reflect different selection pressures on the phage community. For example, podoviruses typically have a much narrower host range than myoviruses (Suttle and Chan, 1993; Waterbury and Valois, 1993; Wichels et al, 1998; Sullivan et al, 2003), although this was not the case in this study. Additionally, many siphoviruses and filamentous phage are temperate, which has further implications on lateral gene transfer and lysogenic conversion, as in the case of CTXO and V. cholerae (Waldor and Mekalanos, 1996). 78 Table 5.3. Characteristics of the VpV strains isolated in this study. Viruses are listed alphabetically, followed by strains isolated from oysters (LHs; Chapter II) which were further characterized in this study. Viruses with two genome sizes indicate two co-isolated viruses within the strains (see Results and Discussion). Genome sizes and dimensions are approximate and have been rounded up to the nearest 1 kb or 5 nm (base-plates and tail-fibres, if any, were not included in the tail lengths). Morphotype refers to updated Bradley groupings as described in Ackermann et al. (1984). A horizontal bar (-) indicates the information is not yet available. Question marks (?) indicate that the strain has a suspected co-isolated podovirus which hasn't yet been confirmed. VpV Strain Isolation Host Type Genome Size (kb) Dimensions (nm) Head Tail Virus Family Morphotype Source 261a Clinical 80 60x110 165 Siphoviridae B2 Fraser River Plume 261b Environmental 64 75 140 Siphoviridae Bl Fraser River Plume 262 Clinical 46 50 15 Podoviridae CI Fraser River Plume 263 , Clinical 46 70 200 Siphoviridae Bl Fraser River Plume 360 Clinical 45 75 225 Siphoviridae Bl Sechelt Inlet 376 Clinical 45 60 150 Siphoviridae Bl Malaspina-Hole 377 Clinical 45 50 150 Siphoviridae Bl Malaspina-Lancelot 384 Clinical 45 50 125 Siphoviridae Bl Malaspina-Okeeover 385 Clinical 45 85 150 Siphoviridae Bl Malaspina-Hole 395 Clinical 106 75 150 Siphoviridae' Bl Howe-Mouth 396 Clinical 78 65x135 150 Siphoviridae B2 Howe-Woodfibre 404 \ Environmental: 64 50x75 115 Siphoviridae B2 Jericho Pier HS1 Environmental 64 65 150 Siphoviridae Bl Howe Sound Mil Environmental 83 60x105 145 Siphoviridae B2 Malaspina Inlet PS1 Environmental 64 65 125 Siphoviridae Bl Pendrell Sound SA1 Environmental: 83 i l l l i l l l l l l l t l - l f l l § f : ; l l l l : l l l l l l l t Salmon Inlet LH1 Clinical 44/120 75/95 20/165 Podo/Sipho Cl/Bl Ladysmith Harbour LH2 Clinical 120 95 160 Siphoviridae Bl Ladysmith Harbour LH3a Clinical 120 65 160 Siphoviridae Bl Ladysmith Harbour LH4a Clinical 44/120 85/75 20/170 Podo/Sipho Cl/Bl Ladysmith Harbour LH4c Clinical 44/120 75/100 20/180 . Podo/Sipho Cl/Bl Ladysmith Harbour LH4f Environmental 122 95 165 ' Siphoviridae Bl Ladysmith Harbour LH5a Clinical 44 75 20 Podoviridae CI Ladysmith Harbour LH5d Clinical 44 70 25 Podoviridae CI Ladysmith Harbour LH5f Environmental 125 - - - - Ladysmith Harbour LH6a Clinical' 44/122 ?/95 7/175 Podo?/Sipho ?/B 1 Ladysmith Harbour LH6c Clinical 44 70 20 Podoviridae CI : Ladysmith Harbour . LH7a Clinical 125 95 205 Siphoviridae Bl Ladysmith Harbour LH8a Clinical 125 90 170 Siphoviridae Bl Ladysmith Harbour Figure 5.2. Representative VpV morphologies. A: siphovirus with base-plate and possible tail-fibres (VpV LH2). B: siphovirus with elongated head (VpV 261a). C: siphovirus with needle-like tail (VpV LH1 [120 kb]). D: podovirus (VpV LH1 [44 kb]). Scale bar is 100 nm in all panels. 80 Even though most VpV strains were siphoviruses, there was considerable diversity in dimensions and genome sizes (Table 5.3). The overall range in capsid and tail lengths were -50-135 nm and -15-225 nm, respectively. Water-column VpVs also had a large variety of genome sizes (-45-106 kb), with 69% of the strains in the <69 kb size-class. Pulse-field gel electrophoresis (PFGE) of virioplankton (Wommack et al, 1999a; Steward et al., 2000) and cultivation studies (Wichels et al, 1998) have shown that 50-90% or more of marine phage belong to this size-class. The genome size appeared to depend on the host used for isolation. Isolates on the environmental host had genome sizes of either 64 or 83 kb, whereas most isolates on the clinical host were 45-46 kb. By comparison, the oyster VpV strains were much less diverse in their genome sizes, and were dominated by 44 kb podoviruses and 120-125 kb siphoviruses. The presence of two different phages in four individual clones (LH1, 4a, 4c, and 6a) was first observed by PFGE and then confirmed via TEM (Figs. 5.2C and 5.2D) for at least three of the four (LH1, 4a, 4c). Precedence exists for clones of VpVs containing two phages. Kellogg et al. (1995) isolated a VpV strain from the coast of Florida that, upon sequencing (Seguritan et al, 2003), revealed two closely related phage. As in the case of Seguritan et al (2003), given the multiple rounds of purification and amplification from single plaques, it seems likely that the phages were co-isolated and co-amplified in a mixed lysate, and it is unlikely that these are helper-satellite virus systems such as P2-P4 (Lindqvist et al, 1993). 5.4.3 VpV Host-Range Diversity The host ranges of the water-column VpVs were determined using V. parahaemolyticus strains from water, oysters, and sediment; as well as other Vibrio spp. (Tables 5.2 and 5.4). The VpVs infected between 4 and 13 strains (mean = 7.4), and every host-range pattern was unique. Half of the viruses were not species-specific and infected two strains of V. alginolyticus. This is uncommon,, as most VpVs infect only V. parahaemolyticus strains (e.g. Koga et al, 1982; Kellogg et al, 1995). Only P4 (Baross et al, 1974) and KVP20 (Matsuzaki et al, 1998) were known to infect other Vibrio spp. (as the VpVs in this study), and only vl4 (Nakanishi et al, 1966) and KVP40 (Matsuzaki et al, 1992) have been reported to infect other genera. This discrepancy is probably real given that there is confidence in the identification of the V. alginolyticus strains. A possible explanation is that the viruses from this study represent all of the podovirus and a significant amount of the siphovirus diversity yet encountered for VpVs, indicating that these broader host-range types simply had not yet been discovered. Overall, the 81 viruses infected more of the oyster and sediment bacteria (55 and 46% of the total pairings, respectively) than bacteria from the water column (<13% of the pairings), which were isolated from the same environment as many of the viruses. Similarly, VpVs isolated from oysters also had lower sensitivity against water-column bacteria (Chapter II). The cause for this resistance is unknown. Abundances of VpVs in the water column were very low (<1 L"1) in this study and accompanying abundances of V. parahaemolyticus are also typically low (<l-70 mL"1) in North American coastal waters (Kaper et al, 1981; Kelly and Stroh, 1988; DePaola et al, 1990). One would anticipate selection for permissive bacteria in this scenario as the contact rates between the viruses and hosts are too low to allow for the selection of co-occurring.resistance (Waterbury and Valois, 1993; Suttle and Chan, 1994). However, low contact rates can lead to selection for lysogeny (reviewed in Wommack and Colwell, 2000), which could prevent some VpVs from infecting some bacteria (super-infection prevention; Stewart and Levin, 1984). The host-range data were converted to a dendrogram (Fig. 5.3) to allow comparison among the VpVs. In general, the water-column viruses were separated from the oyster viruses, with only 2 of 16 water viruses (VpVs 376 and 384) not within the water-column clade (HR1). Although the two environments were separated, there was no strong geographic separation within the water VpVs. Similar locations were dispersed throughout the tree, indicating that the diversity within a site was as high as among sites. The exception was a cluster of 5 VpVs containing the 4 Fraser River Plume (FRP) strains within the HR1 clade. It is possible that the estuarine conditions within the plume favour the selection of a narrower subset of V. parahaemolyticus host strains, which in turn select for a more specific host range among the viruses from that location. There was no clustering associated with whether an environmental or clinical host was used for isolation, or based on the morphology of the viruses. There were, however, some groupings associated with genome size. Within the oyster VpVs (HR2), there were clustered pairs of similar genome sizes, including a double-virus pair (44 / 120 kb), but the pairing was not consistent and did not hold for the water-column VpVs (e.g. 83 kb Mi l and SA1 far apart). Overall, the major determinant of host-range diversity was not geography, but the source environment from which the VpVs originated. 82 Table 5.4. Host range of water-column VpV strains against co-occurring Vibrio parahaemolyticus from three environments and related Vibrio spp. (see Table 5.2). Solid squares indicate lytic infections, whereas open squares indicate the absence of lysis. Viral Strain Host (Species and Strain) < Xt < < < < t l t3 13 t3 < < : < < : < X < < < < X < < X < < X < < X < O v O \ O v O \ O N ^ ] ^ ] 0 0 0 0 v £ > \ O ON —1 H - >-> to U> Ul ON < x < O < X < X 00 < x < < x < X 00 •a < 00 > BP §|1|1 ~. • • , . . liliii - : . lliill lip ii§ Iii • lllpf; Hll H llfpl jjjjjj 1111 Hill iliili 1111 111 MB Wmmk lilllipllill^  111 \ ft ill!! 3 V. parahaemolyticus C00-9a V. parahaemolyticus JOO-5 V. parahaemolyticus J00-6a V. parahaemolyticus J00-7a V. parahaemolyticus J00-9a V. parahaemolyticus S00-1 V. parahaemolyticus SO 1-31 i « V. parahaemolyticus V408-10 V. parahaemolyticus V416-7 V. parahaemolyticus V424-A3 V. parahaemolyticus V439-A1 V. parahaemolyticus V465-A1 V. parahaemolyticus V478-1 V. parahaemolyticus V497-B1 W5 a 3 n s V. parahaemolyticus S00-2 V. parahaemolyticus S00-3 V. parahaemolyticus S00-5 V. parahaemolyticus S00-8 V. parahaemolyticus SO 1-20 m o •a V. alginolyticus ATCC 17749 V. alginolyticus C00-6a V. alginolyticus JOO-lOa V. alginolyticus PWH3a V. alginolyticus S99-45 V. alginolyticus S00-4 V. alginolyticus V478-16 V. alginolyticus V478-17 V. alginolyticus V497-C16 V. fluvialis S99-44 V. natriegens ATCC 14048 V. vulnificus S99-48 V. vulnificus S01-38 83 Host Genome Morph Soma LH6c 44 l l l i i LH 262 r_;4'6':v|:; FRP 385 ::/i>:?45::; l i l t III MI 263 '.;:::|;46;||| l i l l f l l - FRP 261a l l l?< i l l s* FRP 261b l l l l 1 II1IKII l i i l i i , s FRP SA1 | I E | I ;-- :-|83i|| - | l | |s i / 377 1; :.45;|:;| MI LH5f ' E i i i i i i i i i l i l l l l f f LH 360 f i l - ' l f SI HS1 l l i l l l l l M K I | l s: l | I1IHS| 395 :1||I|§I':;: I I s i l l |||HS|. LH6a 44/122 - /s LH MI1 list | | | -83| l l i l i i i l l l l M i l PS1 M i l 64 Bi$ l l : ; l l ? s l 396 78 l l | S ? | | ||::HS| 404 1 1 1 64 ll:-§?: 1:1 JP LH4f E l l l l l l l l l l l l l l l l LH LH4a 44/120 i i i i i i LH 376 | l - | 4 5 | l i i s i - i | | |MI* 384 •;::i|::45i;V::;:: i-illiii? lllii:-LH1 44 / 120* i i i l i i LH LH4c 44/120 P /S LH LH5a 44 . l l l l l l l l LH LH5d 44 p LH LH2 120 1 S m i l LH3a i l l f i j f i i s i i i - p LH7a l||i25f |t | l l l f l l i 111!? LH8a l l l l l l l l 111 s i l l LH HR1 HR2 0.1 Figure 5.3. Phenotypic diversity of VpVs. UPGMA tree of the VpV host-range similarities resulting from 928 crosses (29 VpVs x 32 hosts). Two host-range clades (HR1 and HR2) are indicated. VpV strain designations are those listed in Table 5.3. Also included are which host type the VpVs were isolated upon (E - environmental; clinical otherwise), the genome size (in kb), morphotype (M = myovirus; P = podovirus; S = siphovirus; S* = elongated-head siphovirus), and geographic origin (corresponding to Fig. 5.1). Viruses isolates from oysters are shaded in grey, whereas viruses from the water column are not shaded. The scale bar represents a dissimilarity of 10%, corresponding to a difference of 5.4 infection/non-infection pairs between strains. oo 4^  5.4.4 VpV Genotypic Diversity The genetic diversity of the water-column and oyster VpVs was determined using degenerate-primed random amplification of polymorphic DNA (DP-RAPD; Fig. 5.4). This PCR-based technique allows for whole genome comparisons (strain-typing) of viruses and does not require a priori genetic information (Chapter IV). The DP-RAPD assay yielded reproducible fingerprints that were converted to a similarity dendrogram (Fig. 5.5). Overall, the genetic diversity of the VpVs was high, as evidenced by the multiple, deep-branched groupings. As with host range, the water-column VpVs (clades Wl-3, plus 262 and 396) were separate from the oyster VpVs which clustered in a few groupings of 2 or 4 strains (clades 01-3, plus LH2). However, on a finer scale, individual pairs (or small clusters) of viruses that grouped together by host range were not genetically similar. This is perhaps not surprising given that unrelated viruses from different families (such as T4, T7, and lambda) can infect the same host (such as Escherichia coli). The reference coliphages (T4, T7, and lambda) were well separated (>20% dissimilarity) from their closest vibriophages. There was some separation based on morphology. A cluster of podoviruses, primarily containing the 44 kb LH viruses (clade 03), was grouped with VpV 262 and, even more distantly, with T7. VpV 262 is known to be a distant relative of T7 (Hardies et al, 2003) and its separation from the LH podoviruses suggests that it is also quite different from the oyster podoviruses. There was good clustering, in two pairs (Wl and 02) and one foursome (W3), of the VpVs isolated on the environmental host. The VpVs within these clusters also shared similar genome sizes. This suggests that there was a genetic bias to retrieving some strains from the environment. Interestingly, this selection bias was not seen in the host ranges of the VpVs. Some examples from this study were the Howe Sound strains 395, 396, and HS1. These viruses had similar host ranges (HR1 clade, Fig. 5.3) yet individual enrichments using the clinical host yielded two 78 and 106 kb siphoviruses, whereas the pooled enrichment using the environmental host with the same samples yielded a very different 64 kb siphovirus. 85 "T3 Figure 5.4. VpV DP-RAPD fingerprinting. Gel image showing the results of the DP-RAPD on viruses isolated in this study and additional strains for comparison. Strain designations are those listed in Table 5.3. Coliphages T4 (myovirus), T7 (podovirus), and Lambda (siphovirus) are included for reference. L: 100 bp ladders (100-1,500 bp + 2,000 bp top band). 97.5% 0.1 Host Genome Moiph Source LH2 , 120 l l l s f l l LH T4 -MM E 83 MI SA1 :::V:::::::83?:.:;.:;i. -LH1 44 / 120 LH LH3il 120 s LH LH4.1 44 / 120 • P/S LH LH7a 125 s LH 396 HS LH4C P/S LH LH8.1 | l l l 2 5 l | | s LH 360 45 ins 1 a i l 384 45 385 45 MI ;';:| 377 45 MI 261a FRP 263 FRP 376 LH4f ' . E 122 1 s • LH LH5f E 125 - LH A:';;-;:ri 3''::S|::;1:: LH5n P I. H LH5(I i i l t l t l t P - LH LH6c 44 P LH LH6a ililll LH 262 P T7 ' :f: ;:!::fi:4Q";::% P 395 M:06:i;;::: •::->; sm: i l H S i l HS1 I M I s HS PS1 s -RSlli 2611) s 404 ; |s;E|| iv-ijp i i W1 01 W2 02 031 W3 Figure 5.5. Genotypic diversity of VpVs. UPGMA tree of the VpV genetic fingerprint similarities resulting from the gel image in Fig. 5.4. Three clades containing water-column strains (Wl-3) and three clades containing oyster strains (01-3) are indicated. VpV strain designations are those listed in Table 5.3. Also included are which host type the VpVs were isolated upon (E = environmental; clinical otherwise), the genome size (in kb), morphotype (M = myovirus; P = podovirus; S = siphovirus; S* = elongated-head siphovirus), and geographic origin (corresponding to Fig. 5.1). Viruses isolates from oysters are shaded in grey, whereas viruses from the water column are not shaded. Reference strains (lambda, T4, and T7) are outlined in boxes. The scale bar represents a dissimilarity of 10%. The vertical line is the percent similarity (97.5%) of the identical standards ran during the analysis - therefore, any groupings to the right of the line should be considered identical or very nearly so. 00 Similar genome sizes, potentially indicating near-identical strains of a virus-type, clustered together fairly well. The 64 kb (W3 clade), the 83 kb (Wl), the individual 44 kb (03), most of the 45/46 kb (W2), and most of the >120 kb (within 01, 02) VpV groups were clustered together. Of course, having a similar genome size does not prove relatedness given that the size-ranges of the three tailed-phage families overlap (National Center for Biotechnology Information, 2004) and given recent comparative phage genomic evidence of rampant gene transfer within "local" phage groups or even between distantly related phages types (Hendrix et al, 1999; Pedulla et al, 2003; Hendrix, 2003). However, the two concepts are not mutually exclusive and there is ample evidence, such as from dairy phages (Briissow and Desiere, 2001; Briissow and Hendrix, 2002) and vibriophage (Matsuzaki et al., 2000; Seguritan et al, 2003), of highly related phages (determined via whole genome sequence or DNA/DNA hybridization) sharing nearly identical genome sizes. Single VpV strains with unique genome sizes (78, 80, and 106 kb) branched out more deeply. The double-viruses were not all clustered together, but were within groups matching either their small virus (LH6a; 03) or their larger virus (LH1, 4a, and 4c; 01). Although it is difficult to make assessments regarding relative contributions of templates in the DP-RAPD assay (Chapter TV), it is at least evident that there are some differences among the double-virus strains. Additionally, the clustering of the double-viruses with individual small and large genome-size strains provides further evidence that these systems were the result of co-isolations, and not distinctly different satellite-helper systems that required both viruses for propagation. Finally, genetic diversity of the phage was not tied to geography, but to the environment from which the VpVs originated. Water-column VpVs (e.g. HS1 and PS1 in W3) from locations with very different oceanographic conditions, and separated by up to -120 km, were very closely related. Conversely, some VpVs originating from samples within the same inlets (e.g. MI and SI) were quite different. The only VpV distribution data available for comparison is work by Kellogg et al. (1995) which showed that genetically related VpVs were distributed throughout the coast of Florida and as far away as Hawai'i, indicating that perhaps homogeneity within the world's oceans should be anticipated. Investigations with cultured cyanophages (Wilson et al, 1993) and MpVs (Cottrell and Suttle, 1991), as well as evidence from algal virus DNA polymerase genes (Short and Suttle, 2002), cyanophage structural genes (Zhong et al., 2002), and podovirus polymerases (Breitbart et al, 2004b), have shown the widespread distribution of specific virus-types in the world's oceans. 88 5.4.5 Conclusions Viruses infecting V. parahaemolyticus were found to be widely occurring in the water column of the Strait of Georgia. The assemblage of VpVs characterized in this study substantially increases the known morphological and host-range diversity, and increases the genetic understanding of VpVs. Insights into phage diversity and evolution in general have already been obtained by the complete sequence of one of these phages, the 46 kb podovirus VpV 262 (Hardies et al, 2003), and two more sequences are underway: VpVs 261a and 263 (46 and 80 kb siphoviruses, respectively). Both the host-range and genetic diversity of the VpVs demonstrated separation of water-column strains from related oyster strains. This separation of diversity mirrors their hosts' diversity, as determined by a recent study of Vibrio spp. in coastal British Columbia (Chapter III). The patterns of phenotype (host range) and genotype were, however, uncoupled on a fine scale. In other words, the most similar genotypes did not concurrently generate the most similar host ranges. Overall, the major determinant of host-range and genetic diversity was not geography, but the source environment from which the VpVs originated. Viruses from some of the farthest separated locations (-120 km) were closely related, while the diversity within some locations was as high as between locations. Therefore, VpVs within and adjacent to the Strait of Georgia are a highly diverse, yet homogeneous mixture of phenotypes and genotypes. This study is only the second to look at distribution of VpV genotypes, and the first to obtain whole-genome comparisons of VpVs to host-range diversity within an ecological context. 89 Chapter VI: Conclusions The focus of this dissertation was the examination of the distribution and diversity of viruses infecting a single bacterial species in multiple environments. Members of the Vibrionaceae, which are globally distributed (Thompson et al, 2004a), and Vibrio parahaemolyticus viruses (VpVs) served as a model for heterotrophic host-virus interactions in the marine ecosystem. Vibrio spp. and VpVs were investigated within oysters and sediment, as well as the water column which serves dually as a habitat and a connecting medium between the former two environments. Traditional culture-based approaches were used together with molecular tools, including a newly developed virus strain-typing method, to isolate and characterize the host and virus populations. 6.1 Host-Virus Dynamics within Oysters The first research section of the dissertation (Chapter II) dealt with the host-virus relationship within molluscs. The viruses within the 7,000 t of oyster harvested in BC in 2003 (Fisheries and Oceans Canada, 2004) contain the viral load of the entire volume of the Strait of Georgia (Thomson and Foreman, 1998) seven times over at the usual abundances of VpVs measured in the water column (Chapter V). It is therefore evident why investigating the phage-host dynamics within oysters was important in understanding the ecology of these viruses. It is also important as a model for other Vibrio spp. (and other heterotrophic genera) which are present in the pelagic environment, yet in much higher abundances associated with some sort of strata (e.g. animal commensals, biofilms, sediments). This was the first such study to follow the seasonal dynamics (inside, and outside, of oysters) of Vibrio spp. and the viruses which infect them, combining abundances with data on the host range of viral strains and the efficiency with which individual strains of V. parahaemolyticus recovered infectious viruses from oyster and seawater samples. Together these data provided a comprehensive picture of changes in the host and virus communities over several months. It was evident that two distinct regimes existed. There was a summer regime during which viral production in bacterial populations within oysters was presumed to occur, and a winter regime during which the virus population persisted in the absence of detectable hosts. Although the results emphasized that viruses were significant agents of bacterial mortality within oysters, calculations of the virus-induced mortality on V. parahaemolyticus demonstrated that the presence of additional host species was required to sustain viral production within oysters, possibly even during summer. The fact that the VpVs were capable of infecting other Vibrio spp. demonstrated that this would be possible. Several authors have suggested research of this 91 kind in order to determine the extent of polyvalency (broad host range) and to provide support for the theory that polyvalent phage would be anticipated when/where host cell concentrations are low (Wommack and Colwell, 2000; Chibani-Chennoufi et al, 2004). Further temporal changes in the viral host-range patterns demonstrated, for the first time, that a significant shift in virus strains occurred during the winter months. Winter VpV production was therefore not supported by summer bacterial strains and the system was probably maintained by a dynamic mixture of altered host culturability and viral exploitation of cross-species infection. These findings, along with examples such as seasonal changes in UV resistance among cyanophages (Garza and Suttle, 1998) and strain variation during the progression of an algal bloom (Tomaru et al, 2004b), confirm that host-virus systems respond to changing environmental conditions with changes in their phenotypic composition. Additionally, these results have implications for other pelagic host-virus systems that also have high abundances in other environments and/or that respond to strong seasonal trends. Production of many other marine viruses may depend on inputs from non-water-column sources and contributions from hosts beyond the primary species. 6.2 Host Distribution and Diversity within the Water Column and Sediments The second research section of this dissertation (Chapter III) focused on the hosts of VpVs within the water column and sediments. The polyvalency of the VpVs from oysters in the previous chapter highlighted the necessity of investigating Vibrio spp. in general, not just the primary host V. parahaemolyticus. The purpose of the study was therefore, to determine the extent of Vibrio spp. populations in the two environments along the coast of British Columbia and to determine whether there were geographic trends in their genetic diversity and phage-typing patterns (phage susceptibility). Bacteria initially isolated from oysters (Chapter II) were further characterized in this chapter and used as comparison to the water-column and sediment strains. The ubiquitous distribution of Vibrio spp. in the environments that were sampled suggested that they could play a significant role as hosts for viral infection within the bacterioplankton. Not surprisingly, geographic distribution was a poor determinant of strain diversity. Environment of isolation (water, sediment, or oyster) however, had a much more profound effect on genetic diversity (via ERIC-PCR fingerprinting), and on phage-typing patterns to an even greater extent. Susceptibility patterns of isolated Vibrio spp. demonstrated that sources of high abundance (sediment and oysters) generally produced V. parahaemolyticus strains with high susceptibility to viral infection; and the opposite was true for the water column which produced 92 highly resistant strains. This is the first known investigation of the distribution of V. parahaemolyticus genotypes and phenotypes (phage-types) in multiple environments. The previous chapter showed temporal changes in diversity, whereas these data demonstrated spatial diversity. The results showed that there was segregation of bacterial host strains in different environments, under differing selection pressures, which will have an impact on in situ phage production. The findings, along with the data from Chapter II, may help address the problem of maintaining virus populations infecting host species that are at low abundances in the water column (Wiggins and Alexander, 1985; Wilcox and Fuhrman, 1994; Wommack and Colwell, 2000). Their maintenance may rely on a small amount of water-column production, along with larger inputs from other environments. 6.3 Development of a Method to Assess Viral Diversity Since the previous chapter benefited from the availability of a genetic diversity method, the penultimate research section of the dissertation (Chapter TV) involved the development of a molecular typing tool for the assessment of phage diversity. Most methods that existed, such as DGGE and RFLP of gene products, required a priori genetic information on the viruses in question. However, there was a need to analyze a large collection of potentially unrelated viruses with no known sequence information. Other available methods for examining novel viruses, such as restriction digests of genomic DNA and DNA/DNA hybridizations, were potentially not universally applicable (i.e.: different restriction enzymes for different viruses) and required significant quantities of extracted DNA. This led to the modification of random amplification of polymorphic DNA (RAPD) using a degenerate primer to produce unique and reproducible banding patterns from viral genomes. In the degenerate-primed RAPD analysis (DP-RAPD), a selection of algal virus and bacteriophage strains were profiled that encompassed an array of genome sizes and virus families. Closely related viruses (e.g. strains infecting Micromonas pusilla) generated similar, yet unique DP-RAPD patterns that could be readily distinguished from viruses within the same family (Phycodnaviridae) infecting a Chlorella-like alga. As well, marine vibriophage from the families Myoviridae, Siphoviridae, and Podoviridae showed high diversity and were distinct from coliphage and cyanophage. Results from mixtures of viral templates indicated that the assay would probably be of limited use in profiling complex natural viral communities, unless it complemented existing methods such as PFGE. However, the assay was fairly robust in the f 93 presence of host template contamination and provided an excellent method for strain-typing and assessing the diversity of groups of novel viruses. 6.4 Virus Distribution and Diversity within the Water Column The DP-RAPD assay was then used in the final research section of the dissertation (Chapter V) in which the purpose was to isolate and characterize VpVs from the Strait of Georgia, and to examine their spatial variation in host-range and genetic diversity. Although the high production rates of viruses within oysters, which are continuously shed into the environment, showed that the oysters served as the major environmental source of VpVs (Chapter II); the water column transmits these populations so that they may seed other locations (sediment, zooplankton, molluscs) and seasons. Viruses initially isolated from oysters (Chapter II) were further characterized in this final chapter and used as comparison to the water-column VpVs. Viruses infecting V. parahaemolyticus were found to be a widely occurring component of the virioplankton, but typically at very low abundances (<1 L"1) in the Strait of Georgia region. Contrary to previous isolations, most VpVs were siphoviruses, although podoviruses were also common, and genome-sizes diversity was high (45-106 kb). By comparison, the oyster VpV strains were much less diverse in their genome sizes, and were dominated by 44 kb podoviruses and 120-125 kb siphoviruses. To my knowledge, these are the first podoviruses that have been isolated that infect V. parahaemolyticus. Half of the viruses were not species-specific, uncommon for VpVs, as they infected two strains of V. alginolyticus. Both the host-range and genetic diversity of the VpVs (via DP-RAPD) demonstrated separation of water-column strains from related oyster strains (Chapter II). This separation of diversity based on environment mirrored their hosts' diversity, as described for the Vibrio spp. in Chapter III. The patterns of phenotype (host range) and genotype were, however, uncoupled on a fine scale. In other words, the most similar genotypes did not concurrently generate the most similar host ranges. This study was only the second to look at distribution of VpV genotypes, and the first to obtain whole-genome comparisons of VpVs to host-range diversity within an ecological context. Molecular-based investigations of viral gene diversity have, at times, found evidence for both geographic separation (Zhong et al, 2002; Frederickson et al, 2003) and for the lack of geographic barriers (Short and Suttle, 2002; Breitbart et al., 2004b). It was of interest, therefore, to know whether culture-based isolations of viruses would show segregation of strains in any way. > 94 Overall, the major determinant of host-range and genetic diversity was not geography, but the source environment from which the VpVs originated. Viruses from some of the farthest separated locations (-120 km) were closely related, whiie the diversity within some locations was as high as between locations. Therefore, VpVs within and adjacent to the Strait of Georgia are a highly diverse, yet homogeneous mixture of phenotypes and genotypes in the water column, distinct from their counterparts within oysters. 6.5 Conclusions and Contributions Overall, the diversity of viruses infecting a single heterotrophic species was high and the host-virus populations were under similar phenotypic and genotypic selective pressures, mediated by the source environment and not geography. The assemblage of VpVs characterized in this study substantially increases the known morphological and host-range diversity, and increases the genetic understanding of VpVs. When compared to all 112 previously described VpVs (summarized in Table 6.1), the 33 (double-virus strains as 2 each) virus strains from this study now represent almost 25% of the total known diversity. Additionally, they represent more than half of all siphoviruses, are the only podoviruses so far isolated, and (with the exception of P4) comprise all of the VpVs from shellfish. Of interest to those working with PFGE and genome evolution, this pool of viruses provides a large increase (200% for VpVs) in the known genome sizes from cultured marine heterotrophic bacteriophage. Significant insights into phage diversity and evolution have already been obtained by the complete sequence of one of these phages, the 46 kb podovirus VpV 262 (Hardies et al, 2003), which became only the third sequenced marine phage. Two more sequences are underway - VpVs 261a and 263 (46 and 80 kb siphoviruses, respectively) - by a group at the Pittsburgh Phage Biology Institute interested in the differential photorepair capabilities (data not shown) between these two viruses. Yet another group at the Massachusetts Institute of Technology is interested in the distribution and diversity of VpV 262-like genes in the environment. These examples show that these cultured viruses, along with the conclusions of this dissertation, have already begun, and hopefully will continue, to provide the impetus for new investigations into marine phage ecology. 95 Table 6.1. Chronological listing of all characterized Vibrio parahaemolyticus Viruses (VpVs) in the literature. Viruses were included in this list if they were isolated upon (or primarily infected) V. parahaemolyticus and were functional phage isolates. Morphotype refers to updated Bradley groupings as described in Ackermann et al. (1984). Host range is described as being strain-specific ("Strain"), infecting multiple V. parahaemolyticus strains ("Vp"), infecting the host species + other Vibrio spp. ("Vibrios"), or as being a broad host-range phage ("Broad") capable of infecting species in at least one other genus. Viruses are listed by year first isolated/described, and publications with later dates indicate further characterization and/or sequencing of the virus. VpV Strain Family Morpho-type Genome Size (kb) Host Range Source Year References Characterized v6 Inoviridae Vp Culture - Spontaneous vl2 Vp . Japanese Patient 1966 Nakanishi etal., 1966 vl4 Broad Coastal Japan nonea Myo-/Siphoviridae Al/Bl Vp Coastal NC Sediment 1973 Sklarow et al., 1973 P4 Myo-/Siphoviridae AI/UI Vibrios Japanese Shellfish 1974 Baross etal, 1974 Vpl5P Myoviridae Al Vp Coastal Japan Vpl7P Siphoviridae Bl Strain Coastal Japan 1978 Hidaka and Tokushige, 1978 Vp25P Myoviridae Al Vp Coastal Japan Vp33P Siphoviridae Bl Vp Coastal Japan VP3 Siphoviridae Bl 65 Vp Coastal Japan 1981 Koga and Kawata; 1981; Koga and Kawata, VP1 Myoviridae Al Vp Coastal Japan + 9 other Myoviridae Al Vp Coastal Japan VP5 Siphoviridae B2 Vp Coastal Japan 1982 Kogaefa/., 1982; + 3 other Siphoviridae B2 Vp Coastal Japan Koga and Kawata, 1991 VP6 Siphoviridae Bl 74 Vp Coastal Japan + 2 other Siphoviridae B1 Vp Coastal Japan TP1 Siphoviridae Bl 65 Vp Culture - Induced 1991 Koga and Kawata, 1991 VP143 Myoviridae Al 48 Vp Culture - Induced 1991 Muramatsu and Matsumoto, 1991 VP253 Myoviridae Al 48 Vp Culture - Induced OPEL8C-1 Myoviridae Al Strain Coastal Florida + 31 other (DHAW1-5 Myoviridae Myoviridae Al Al Strain Strain Coastal Florida Coastal Hawaii 1995 Kellogg etal, 1995 ' + 33 other Myoviridae Al Strain Coastal Hawaii KVP20 Myoviridae A2 250 Vibrios Coastal Japan 1998 Matsuzaki etal, 1998 lvpf5 Inoviridae l l l i - r l i i : ! 8.5 Vp Laos Patient 1999 Nakasone etal, 1999 KVP241 Myoviridae Al 130 Vp Coastal Japan 2000 Matsuzaki et al., 2000 + 6 other Myoviridae Al Vp Coastal Japan ON Table 6.1. Continued. VpV Strain Family Morpho-type Genome Size (kb) Host Range Source Year References Characterized and Sequenced Vfl2 Inoviridae 7.965 Strain C^ulture - Spontaneous 19X4 Taniguchi etal.. 1984; Vf33 lnoviridae 7.965 Strain Culture - Spontaneous Chang etal., 1998 KVP40 Myoviridae A2 245 Broad Coastal Japan 1992 Matsuzaki etal., 1992; Miller etal., 2003 VP16Tb Siphoviridae Bl 49.6 Strain Coastal Florida 1995 Kellogg etal, 1995; VP16Cb Siphoviridae Bl 47.5 Strain Coastal Florida Seguritan etal, 2003 Vf03K6c Inoviridae - 8.782 Strain Culture - Innate 2000 Nasu etal., 2000 Vf04K68 lnoviridae ttlfp|l|t§l 6.891 ; v P Culture - Innate 2002 Change?al, 2002 VpV 262d Podoviridae CI 46 Vibrios Coastal British Columbia 2003 Hardies et al, 2003 a This viral isolate was not given a designation by the authors. b VP16T and VP16C were initially published as the single phage 016. 0 VT03K6 is also known as filamentous phage f237. d VpV 262 is the same phage as presented in Chapters IV and V. 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